BACTERIAL COMMUNITY PROFILING OF WESTERN AUSTRALIAN BOBTAIL LIZARD (TILIQUA RUGOSA)

Ruby Mckenna

Bachelor of Science

Submitted in fulfilment of the requirements for the degree of Honours in Biological Sciences

School of Veterinary and Life Sciences Murdoch University, Perth 2019

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Statement of Original Authorship

I declare that this thesis is my own account of my research and contains as its main content work which has not previously been submitted for a degree at any tertiary education institution.

Ruby McKenna

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Abstract

Ticks are haematophagous and major vectors of pathogenic microorganisms. In

Australia, over 74 species of ticks have been described, of which 13 are known to parasitise reptiles. While only three borne diseases are formally recognised, honei, the causative agent of Flinders Island , has been long associated with the reptile tick,

Bothriocroton hydrosauri, despite this tick rarely reported to parasitise people. More recently, a novel Rickettsia species, Rickettsia gravesii, was reported in the ornate kangaroo tick (a common human biting tick), triguttatum, on Barrow Island in Western Australia.

In addition, a number of overseas tick-associated microbes (taxa of interest) have been identified in Australian human-biting ticks using an advanced molecular technique, next generation sequencing (NGS). Therefore, the aims of this study were to morphologically and molecularly identify ticks that parasitise reptiles, specifically the bobtail lizard, Tiliqua rugosa, and to employ NGS to profile the bacterial 16S rRNA gene (16S) within the ticks. The taxa identified would then be phylogenetically compared to known taxa of interest.

A total of 306 ticks from Western Australia were morphologically identified and 30 were removed from the data set and the remaining 276 included all developmental stages comprising males 43.1% (n=119); nymphs 40.6% (n=112); females 15.2% (n=42) and three larvae (1.1%). Using Australian-specific morphological keys, ticks were identified as A. albolimbatum. To provide more accurate species identification, molecular barcoding of the cytochrome C oxidase 1 (CO1) gene was employed on 17 nymphal and two larval ticks, along with Northern and Southern WA A. albolimbatum ticks as representative specimens. However, only one nymph (5.8%) and two larvae (100%) generated clean chromatograms and all were identified as A. albolimbatum; the two larvae were genetically identical to the Southern representative sample and the nymph more genetically similar to the Northern representative iii

sample (0.48% genetic difference). The genetic distance between the two A. albolimbatum sequences was 4.79%.

A total of 116 ticks and six controls were processed for 16S metabarcoding to profile the bacterial communities. A total of 9,706,920 reads were generated using an Illumina V3 600 cycle run on the MiSeq platform. A final quality filtered data set consisted of 4,823,227 reads, with a total of 1,385 zero operational taxonomic units (ZOTUs) generated. The bacterial diversity of the ticks was observed to be statistically different between life stage, with males exhibiting the highest diversity. The bacterial microbiome of the tick samples was dominated by the phylum (90.94%) and also included Actinobacteria (3.81%) and

Firmicutes (3.71%). Interestingly, orders and , which contain known taxa of interest, were identified. BLAST analysis of ZOTUs associated with taxa of interest revealed the most abundant ZOTU as Rickettsia endosymbiont (100% similarity;

GenBank accession MK00580); ZOTU2 was 99% genetically similar to Francisella hispaniensis (GenBank accession KT28184); while ZOTU10 and ZOTU19 were most closely related to a Spotted Fever (99.7% similarity; GenBank accession MK30454) and (100% similarity; GenBank accession LC464975), respectively.

Importantly, while R. raoultii does not exist in Australia, literature supports that species delimitation for Rickettsia cannot be based on 16S alone and requires a multi-loci approach.

Therefore, the Rickettsia identified in this study may in fact represent a novel endemic species.

However, the presence of C. burnetii is intriguing and represents the first evidence within A. albolimbatum ticks supported by phylogenetic analysis. While ticks are rarely implemented in the zoonotic transmission of C burnetii, further research is required to investigate the vector competency of this tick and to determine if reptiles, particularly T. rugosa, are viable reservoirs for this bacterium and determine if reptiles pose a risk to wildlife carers.

Overall, this study provides new molecular data for A. albolimbatum and requires further research to investigate the validity of the Northern and Southern morphotypes identified.

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Furthermore, for the first time in Australia, this study presents the bacterial communities within the bobtail tick, A. albolimbatum, and showcases a rich diversity of microbes, including endosymbionts and known tick-associated pathogens.

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Acknowledgements

Firstly, thank you to my supervisors, Dr Charlotte Oskam (Primary) and Professor Peter Irwin

(Co-supervisor). Charlotte, thank you for your generosity, inspiring ideas and approachable nature.

Thank you to the Vector and Waterborne Pathogens Research Group you have been wonderful.

A special thanks goes to to Dr Jill Austen, Megan Evans and Siobhon Egan. Thank you to

Siobhon, for your mentorship throughout my project. You have exceptionally generous with your time and patience. I would also like to thank Samuel Bollard, Aimee Carpenter, Wenna

Cung for your support throughout this honours year.

Thank you to the many people who contributed ticks to this project and especially to Kanyana

Wildlife Centre for being so helpful with their patient information. I would also like to thank

Gerrut Norval of Flinders University for his helpful tick identification information.

I would like to thank The Harry Butler Institute, The Myrtle AB Lamb Scholarship and the

Loneragan Family Scholarship for their interest in my research, and their generosity in awarding me scholarships.

Finally, thank you to my family and especially my husband Daniel, you are an amazing support.

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Table of Contents

Statement of Original Authorship ...... iv

Abstract ...... iii

Acknowledgements...... vi

Table of Contents ...... vii

List of Figures ...... viii

List of Tables ...... ix

List of Abbreviations ...... x

Chapter 1: Introduction ...... 1

1.1 Tiliqua rugosa (Bobtail) ...... 1

1.2 Ticks ...... 2

1.3 The tick microbiome and tickborne disease ...... 9

1.4 Characterisation of the tick microbiome ...... 11

1.5 Microbes Associated with Bobtails ...... 17

Chapter 2: Materials and Methods ...... 21

Chapter 3: Results ...... 32

3.1 Taxonomic Identification of Tiliqua rugosa ticks ...... 32

3.2 Next Generation Sequencing data exploration ...... 44

Chapter 4: Discussion ...... 63

4.1 Tick identification ...... 63

4.2 NGS data exploration...... 66

4.3 Conclusions ...... 76

References ...... 79

Appendices ...... 87

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List of Figures

Figure 1.1 Phylogeny of the Families, subfamilies and genera of the suborder Ixodida 4 Figure 3.1 Geographic distribution of ticks collected for this study 26 Figure 3.2 Amblyomma nymphal ticks with varying degrees of engorgement 27 Figure 3.3 Female Amblyomma albolimbatum (R76) 28 Figure 3.4 Male Amblyomma albolimbatum (PI1872) with iridescent ornamentation on the scutum 29 Figure 3.5 Amblyomma larval ticks 30 Figure 3.6 Maximum likelihood phylogenetic tree based on 425bp CO1 alignment 33 Figure 3.7 The rarefication curves plotted by the faith phylogenetic diversity metric 36 Figure 3.8 Alpha diversity plots 38 Figure 3.9 PCoA analysis showing the differences in beta-diversity between tick instars 39 Figure 3.10 The abundance of phyla in the Amblyomma albolimbatum ticks and the control samples 40 Figure 3.11. Family level taxonomic assignment for each instar of the tick Amblyomma albolimbatum 40 Figure 3.12. Heat map denoting the top 50 genera via relative abundance in each instar of the tick Amblyomma albolimbatum 41 Figure 3.12 Boxplots denoting the percentage relative abundance of the top two genera Francisella and Rickettsia in each instar of the tick Amblyomma albolibatum 42 Figure 3.13 Phylogenetic analysis of 404bp 16S rRNA gene of Rickettsia 44 Figure 3.14 Phylogenetic analysis of 431bp 16S rRNA gene of Francisella 46 Figure 3.15 Phylogenetic analysis of 431bp 16S rRNA gene of the Coxiella-like species 49

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List of Tables

Table 3.1 Instar of ticks and species morphologically identified from Western Australian Tiliqua rugosa. Those designated for DNA extraction in brackets...... 38

Table 3.2 The instar and status of the ticks that were morphologically identified and the number of those ticks which were chosen for DNA extraction for molecular analyses...... 39

Table 3.3 The percentage pairwise distance between all samples sequenced which make up the monophyletic Amblyomma albolimbatum clade. The Reference samples are denoted by “North”, “South”...... 43

Table 3.4 The percentage interspecific differences, with standard error, between the Australian Amblyomma species cytochrome oxidase genes available on NCBI database and those Amblyomma albolimbatum samples sequenced in this study ...... 44

Table 3.5 The samples which had reads for ZOTU 19 (Coxiella) shown with their percent relative abundance from each sample and the relevant metadata...... 60

Table 3.6 Microbial taxa of interest observed in tick samples at a high abundance...... 61

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List of Abbreviations

Abbreviations Disambiguation AGRF Australian Genome Research Facility BIC Bayesian Information Criterion BLAST Basic local alignment search tool bp Base pairs (of nucleotides) DNA Deoxyribonucleic acid dNTP Deoxynucleotide triphosphate DNA EtOH Ethanol gDNA Genomic DNA hr Hour (time) MEGA Molecular Evolutionary Genetic Analysis software

MgCl2 Magnesium chloride

min Minute mM Millimolar mt Mitochondrial NaCl Sodium chloride NCBI Nation Centre for Biotechnology Information PBS Phosphate-buffered saline PCR Polymerase chain reaction RNA Ribonucleic acid rpm Revolutions per min rRNA Ribosomal RNA S Second sp./spp./ssp. Species singular/plural/subspecies Taq Thermus aquaticus DNA polymerase TBD Tick-borne disease

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Chapter 1: Introduction

Herein this literature review, five main areas will be discussed. The first section of this literature review will briefly discuss the lizard Tiliqua rugosa, commonly called the bobtail.

The second section will encompass tick , morphological and molecular identification, and the tick life cycle. Specific attention is made concerning the ticks of the

Australian lizard, T. rugosa. Thirdly, the principle molecular tools and sequencing technologies used to profile will be evaluated. The fourth section will encompass bacterial tick-borne diseases (TBD), common endosymbionts of ticks and the factors affecting tick microbiome. Additionally, bacterial TBD in Australia will be reviewed with specific reference to their aetiological agents (, Rickettsia gravesii, and Coxiella burnetii) with concise synopses of their taxonomy. The microbes associated with bobtails will be mentioned and their potential to act as a reservoir for zoonotic pathogens is discussed. Lastly, the aims and objectives of this study will be outlined.

1.1 TILIQUA RUGOSA (BOBTAIL)

Tiliqua rugosa is an Australian scincid lizard commonly known as the bobtail; however, it is also known as the sleepy lizard, the pine cone lizard and the stump tail lizard throughout different regions of Australia (Wilson and Swan, 2017). The bobtail is a large member of the family Scinidiae, approximately 28-35cm from snout to vent length (ref). The adults are monogamous and have an expected lifespan of 20 to 30 years (Bull, 1988). The females produce 2-3 live young in late summer, with small, stable, overlapping home ranges of 200–

1000 m2 (Bull and Baghurst, 1998).

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This largely herbivorous skink has been found across southern Australia in open habitats that include Mallee woodlands, shrub lands and coastal dunes(Lancaster et al., 2012, Wilson and

Swan, 2017). It is frequently seen in the Perth urban locality, occurring in the pockets of bushland within the inner suburbs (Norval et al., 2019). There are four subspecies of T. rugosa with varying colouration, size and geographic range. The subspecies commonly observed in and around Perth is T. r. rugosa, which occupies the South West of Australia. This subspecies has a stumped tail and its back is variable in the colours brown, olive or black, and also displays obscure to distinct pale bands, with their head coloured orange (Wilson and Swan 2017).

The bobtail is one of the best studied lizard species in Australia, due to the long-term studies that were undertaken by the late Professor C. Michael Bull in South Australia, where it is primarily known as the sleepy lizard. This research was into the parapatric boundary of the hard ticks (Family: ) Amblyomma limbatum and Bothriocroton hydrosauri that parasitise these lizards (Godfrey and Gardner, 2017). In addition to the aforementioned hard ticks, an additional four species of hard ticks and one species of soft tick (Family: Argasidae) have been recorded as parasites of sleepy lizards.

1.2 TICKS

The three main bobtail ticks are the southern reptile tick, Bothriocroton hydrosauri, the reptile tick, Amblyomma limbatum, and the stump-tail lizard tick, Amblyomma albolimbatum (Barker et al., 2014; Smyth, 1973).All of these tick species are found on a variety of reptiles, although they are most frequently found parasitising bobtails, their primary host(Barker and Walker,

2014; Sharrad and King, 1981) Ticks are reported as being one, two, or three host obligate hematophagous arthropods (Jongejan and Uilenberg, 2004; Parola and Raoult, 2001). Three host ticks require a single blood meal at each stage of life stage, and feed on a new host (or reinfect the same host) at each stage in order to complete their life cycle(Parola and Raoult,

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2001; Smyth, 1973). All ticks associated with the bobtail have been documented as three host ticks, excluding the Argasid (soft) tick, Ornithodoros gurneyi.

Ticks are considered parasites because of their reliance on a host organism to obtain sustenance, the blood meal (Jongejan and Uilenberg, 2004). The negative effects by which a tick affects the host ismultifaceted. Firstly, the mechanism by which ticks attach to their host damages the host tissues and their feeding mechanism of alternating periods of sucking blood and salivation which can lead to infection through the insertion of foreign particles and organisms (see section 1.3).Another consequence of the tick attachment is the possible stimulation of the host’s immune system. This stimulation can cause an inflammatory response by eschar or allergic reactions to the incidental saliva secretion, which contains anaesthetic properties (Graves and Stenos, 2017; Jongejan and Uilenberg, 2004). A final consequence of tick attachment is the direct effect on the host’s behaviour, and an indirect effect on the host’s mating partner. These effects are exemplified in bobtails, for they have been observed to move less with a large tick infestation (Main and Bull, 2000), and reports have observed female bobtails leaving their mating partner if they have a large tick infestation (Bull and Burzacott,

2006).

1.2.1 Tick Taxonomy.

Ticks belong to the largest and most diverse phylum: Arthropoda. This phylum represents approximately 80% of all known and is present in almost every region on earth (Parola and Raoult, 2001). Within this phylum, ticks belong to the subphylum and class

Arachnida, along with mites, scorpions and spiders. Ticks and mites belong to the same subclass, , however ticks are further classified into the order Ixodida (Fig 1.1). There are two major extant families of ticks; the Ixodidae, commonly known as hard ticks, and

Argasidae, which are soft ticks. There is also a minor family of ticks, Nuttalliellidae, which consists of a single genus endemic to South Africa (Barker and Murrell, 2004)

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The genera of ticks that belong to the family Ixodidae consist of: Amblyomma,

Anomalohimalya, Bothriocroton, Cosmiomma, , Haemaphysalis, Hyalomma,

Ixodes, Margaropus, Nosomma, Rhipicentor and Rhipicephalus (Fig 1.1;Barker and Walker,

2014). While argasid ticks comprise of only four genera: Argas, Carios, Ornithodoros and

Otobius (Barker and Walker, 2014).

There are approximately 850-900 species of ticks and the current records show that Australia holds 74 of these species(Ash et al., 2017; Barker et al., 2014; Heath and Palma, 2017; Kwak et al., 2018). Of these 74 species, 69 are native, while five species of tick were introduced over

230 years ago when Europeans settled in Australia (Barker et al., 2014). Currently there are

14 soft ticks in Australia, from the two genera of Argas and Ornithodoros. Additionally, there are 60 hard ticks from the genera Amblyomma, Bothriocroton, Haemaphysalis, Ixodes, and

Rhipicephalus.

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Figure 1.1 Phylogeny of the Families, subfamilies and genera of the suborder Ixodida proposed by Barker and Murrell (2004) which has since been amended: The genus Boophilus is now considered a subgenus of Rhipicephalus.

The most significant work on Australian ticks was conducted by F.H.S Roberts (Roberts,

1970), which culminated in the seminal book, ‘Ticks of Australia’. This book created taxonomic keys to identify known ticks of Australia. This set the precedence for tick identification scholarship, and is still used today, as a primary source for identification of wildlife ticks(Barker and Walker, 2014).More recently, a new “Ticks of Australia” book was published by Barker and Walker (2014), which adds a greater depth to tick identification by the use of an image based glossary and precise captioning. This guide, however, is selective and includes only ticks which were considered of importance to the health of domestic animals and humans. Wildlife ticks are not covered, with exception to the southern reptile tick,

Bothriocroton hydrosauri, because it has been known to feed on humans and is regarded as the vector of the causative agent of Flinders Island spotted fever that affects humans (see section 1.4.4).

In addition, there have been two other major developments to the documentation of Australian ticks since Roberts’ seminal work was published. One is the genus Boophilus was demoted to a sub-genus of Rhipicephalus (see Fig 1.1) and the genus Aponomma was dissolved to form the genera Bothriocroton and Amblyomma (Kaufman, 1972; Klompen et al., 2002). The genus

Aponomma was first described by Newmann in 1899 (Kaufman, 1972) as being eyeless and parasitising reptiles. After this first description, all ticks without eyes and found on reptiles were placed into this genus. This was prior to Kauffman (1972), who re-evaluated the genus and described three categories; the typical Aponomma, the indigenous Australian species, and the ‘primitive’ species. Kauffman then further hypothesized that the latter two may represent a new genus. Since then, the indigenous Australian species have been moved to the genus

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Bothriocroton (Klompen et al., 2002), whilst the remaining species moved into the formally well-established genus Amblyomma. Two species that were taxonomically reassigned were the reptile ticks, Amblyomma fimbriatum and Amblyomma trimaculatum (Klompen et al., 2002).

Taxonomic information on wildlife ticks has been somewhat neglected in recent years because they are considered of little medical and veterinary importance. However, studies into parasite infections in wildlife and their roles in ecosystems need appropriate identification so they can be used for proactive measures in managing existing and emerging diseases (Spratt, 2005)

From this, several new identifications of wildlife-associated ticks have been identified in recent years (Ash et al., 2017; Heath and Palma, 2017; Kwak et al., 2018). The ticks of T. rugosa are commonly surveyed in South Australia, however there is a lack of recent data on those in Western Australia (Bull et al., 1984; Norval et al., 2019; Sharrad and King, 1981).

1.2.2 Tick Life Cycle.

The life cycle consists of three instars: larvae, nymph and a sexually mature adult stage

(Jongejan and Uilenberg, 2004; Parola and Raoult, 2001)All instars can be found concurrently on a single host, although moulting to the next life stage takes place off the host (Keirans et al., 1996; Roberts, 1970). The ticks associated with the bobtail are endophilous, nidicolous tick species, and live within the general host environment in nest or burrow (Chilton and Bull,

1993; Kerr and Bull, 2006; T. Petney et al., 1983). Because of their limited mobility, a newly moulted, unfed tick will not advance on a bobtail and will instead rely on the bobtail’s movement for contact (Petney et al., 1983). Evidence suggests that when bobtails share refuges of the same cool temperature that mammals burrow in during high external temperatures, tick burden increases (Kerr and Bull, 2006; Leu et al., 2010). Once a tick is attached to a host, each life stage will remain attached for a period of five days to over two weeks (Chilton and Bull,

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1991; Sharrad, 1979). During this time, ticks will imbibe a large, slow blood meal aided by their extensible cuticle (see section 1.2.3) (Sharrad, 1979). Male ticks do not engorge, although they do take small blood meals, detach and walk around on the host for months while waiting for a female to mate with (Andrews and Bull, 1980)

1.2.3 Tick identification using morphological characteristics

The morphology of ticks is based on the descriptions of two distinct regions of the tick’s body.

The first region is called the idiosoma, which has no abdominal segmentation, and the second region is called the head, which houses the mouth parts called the capitulum, or gnathosoma

(Parola and Raoult, 2001; Sonenshine & Roe, 2013). The entire body and appendages are covered by a chitinous carapace and the idiosoma region contains the eight appendages, the eyes (if present), and the genital aperture (Sonenshine & Roe, 2013). The capitulum is a major morphological feature for differentiating soft ticks from hard ticks. The capitulum is clearly visible in hard ticks as it extends dorsally, while it is less visible in the soft ticks because of its ventral position (Sonenshine & Roe, 2013).

The discrete stage of the lifecycle, which is called an instar, determines morphological characteristics of these two regions (Barker et al., 2014; Sonenshine & Roe, 2013). These morphological identified key characteristics given in the literature are described as: adult ticks display sexual dimorphism with the females being larger than the males and display porose areas on the basis capitulum (Barker et al., 2014 Roberts, 1970; Sonenshine & Roe, 2013).

These porose areas are depressed regions of the cuticle which are perforated with pores.

Females also have a reduced shield, the scutum, which covers the anterior portion of the idiosoma. The larvae and nymphs reflect the adult females scutum, however their scutum will differ in its shape, punctations and ornamentation. The main trait used to separate nymphs and larvae from adults is the absence of a genital aperture and that larvae are hexapod.

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Although the above morphological features are used for identification of instar, they can also be used to identify ticks to the genus and species level. In addition, there are several other morphological features which allow for the identification to the genus and species level, including: the shape of the basal portion and size of the capitulum, the length of the palpal articles and hypostome, the presence or absence of eyes, shape of the spiracular plate, the amount and relative size of spurs on the coxae, the shape of the anal groove, and the presence or absence and number of festoons(Barker and Walker, 2014; Kaufman, 1972; Roberts, 1970).

Another routinely used characteristic is the presence and degree of ornamentation on the dorsal side of the tick, particularly on the adult female. However, this is regarded as an unreliable trait if used as the sole morphological feature because it can vary considerably within species and is often lost in preservation (Parola and Raoult, 2001; Schachat et al., 2018)

Reptile ticks, specifically the genus Amblyomma, have been stated to be hard to differentiate because of their minute differences, and of their morphologically ambiguous characteristics

(Keirans et al., 1996, 1994; Sharrad, 1979).Varying levels of intraspecific ornamentation has been previously observed when surveying reptile ticks in Western Australia(Sharrad and King,

1981). Their study also found a number of discrepancies between tick sizes described in “Ticks of Australia” (Sharrad and King, 1981). Furthermore, there have been theories of hybridisation between A. limbatum and A. albolimbatum because of intermediate species observations

(Sharrad and King, 1981). Although reproductive interference has been researched extensively among the ticks A. limbatum, A. albolimbatum and B.hydrosauri (Andrews et al., 1982) it is out of the scope of this review. However, a close resemblance between A. limbatum, A. albolimbatum, A. moreliae and A. vikkirri tick species, now warrants the revision of the genus

Amblyomma, especially those that parasitise reptiles, using morphological and molecular methods.

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1.2.4 Tick identification through molecular barcoding

Molecular barcoding of a species involves the use of short DNA sequences which are conserved within species and vary between species, in order to identify organisms in an efficient manner with more accuracy than traditional morphological methods (Hebert et al.,

2003). Molecular barcoding has been used as a tool to determine species in the phylum

Arthropoda and has resolved some previously identified cryptic species (Hebert et al., 2003;

Song et al., 2011). Targeting gene loci of the mitochondrial genome has been proven to be more informative than loci on the nuclear genome because of its lack of introns, its limited exposure to recombination and its haploid mode of inheritance (Hebert et al., 2003)

The mitochondrial gene, Cytochrome C Oxidase subunit 1 (CO1), has previously been stated as the standard barcoding gene for phyla (Hebert et al., 2003), and Evans (2019) indicated that it was efficient for barcoding Australian Ixodidae. Because this gene has been used in tick molecular phylogenetics studies, there are sequences available for a limited number of Australian tick species (Barker and Murrell, 2004). Most importantly, for this study, there are CO1 sequences on GenBank for the common bobtail ticks, A. limbatum and B. hydrosauri (Guzinski, n.d.), however there is no sequence available for A. albolimbatum.

Therefore, it will be necessary to sequence the A. albolimbatum CO1 for comparison. One limitation of using a mitochondrial based identification system, is species boundaries can become blurred by hybridisation or introgression. Thus, supplemental analyses of one or more nuclear genes are required (Hebert et al., 2003) .

1.3 THE TICK MICROBIOME AND TICKBORNE DISEASE

Worldwide, ticks are important vectors of infectious agents and are second only to mosquitoes

(Dantas-Torres et al., 2013). This is because of their hematophagous life cycle, which enables

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them to transmit a wide variety of pathogenic protozoa, viruses and bacteria to humans, domestic animals, livestock and wildlife (Jongejan and Uilenberg, 2004; Parola and Raoult,

2001). In Australia, there is an increasing incidence of tick anaphylaxis, mammalian meat allergies and symptoms that are similar to the Northern Hemisphere Lyme borreliosis. In order to understand the possible cause(s), mapping of the tick microbiome, which is a community of protozoa, fungi, viruses, archaea and eubacteria that can be commensal, symbiotic or pathogenic to the host, has been undertaken by several Australian research groups (Egan et al.,

2019; Gofton et al., 2015a; Greay et al., 2018; Loh et al., 2017; Panetta et al., 2017). The current research focuses on the bacterial microbiome of the tick, and it has now been established that both ticks and tickborne pathogens (TBP) are frequently involved in interactions with non- (Bonnet et al., 2017). These non-pathogenic bacteria are likely endosymbionts which a tick may rely upon in order to carry out digestion, reproduction and molting (Bonnet et al., 2017). The following section will discuss the factors which affect the tick bacterial microbiome and the bacterial tick-borne diseases (TBD) documented in Australia.

The bacterial microbiome of an individual tick is thought to be unique due to the factors of species, environment, instar and engorgement status (Bonnet et al., 2017; Parola and Raoult,

2001). Cowdry (1925) was the first to describe species as a factor which affects the tick microbiome. In their study which employed light microscopy, they recognised a difference between Rickettsia like endosymbionts among unfed larvae of different tick species. The environment has also been documented as a determining factor in the tick bacterial microbiome. In an experiment carried out by Zolnik et al. (2016) on ticks were reared in a laboratory setting and their microbiome was compared with those that were collected from the natural environment. They documented that laboratory reared ticks had significantly lower levels of bacterial diversity than the ticks occurring naturally in the environment (Zolnik et al., 2016).

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More recent studies have reported the bacterial diversity is associated with the tick’s life stage

(Lalzar et al., 2012; Thapa et al., 2018; Zolnik et al., 2016). In both Egan (2017) and Zolnik et al. (2016), nymphs were demonstrated to have the highest bacterial diversity when compared to the adult and larval stage. In a study conducted by Lalzar (2012) on the tick species

Rhipicephalus turanicus, Rickettsia comprised almost 10% of the total sequences obtained, and in Rickettsia-positive samples, the relative abundance was higher in males (up to 50%) than in females (up to 10%).

1.4 CHARACTERISATION OF THE TICK MICROBIOME

There has been a substantial increase in the availability of methods, to investigate the tick bacterial microbiome, in the last 40 years. This substantial increase is due to new technological advances regarding the characterisations of emerging infectious diseases, and whole microbiome investigations. Before these tools were readily available, culturing and microscopy of bacteria were the only methods available for detection and characterisation.

This left a lot of infectious diseases to have unknown origins, because many of the infectious agents fell outside the scope of traditional methods of detection. The tools enabling novel characterisations of bacteria are attributed to the sequencing of genetic components and profiling characteristics at the molecular level (Houpikian and Raoult, 2002) These include polymerase chain reaction (PCR) analysis (Mullis and Faloona, 1987), Sanger sequencing

(Sanger et al., 1977), and next generation sequencing (NGS) technologies (Anderson and

Schrijver, 2010).

1.4.1 Polymerase chain reaction (PCR)

PCR is a highly sensitive technology, which has the ability to detect unculturable and low quantity organisms, such as intracellular bacteria (Houpikian and Raoult, 2002). The

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sensitivity of the PCR has enabled data to be obtained from small quantities of DNA and within a reduced timeframe compared to traditional methods (Hopwood, 1999). Moreover, this sensitivity has detected organisms of the tick microbiome, such as Rickettsia (Stenos et al.,

2005), and identified diseases in which the causative organism’s quantity is low in culture from tissue or body fluid (Fredericks and Relman, 1996). While the culture method is seen as traditional and, in comparison with PCR, is more time and fiscally intensive; it still serves as a valuable tool when used in conjunction with the PCR method. For example, the causative agent of Flinders Island spotted fever (R. honei) was first isolated by culture, and was then confirmed by PCR, thus making it further an invaluable tool in identifying morphologically similar organisms (Stenos et al., 2003a).

1.4.2 Sanger sequencing.

The sequencing method eponymously named Sanger or first-generation sequencing, uses dye labelled, chain-terminating dideoxy nucleotides and the enzyme DNA polymerase to create

DNA fragments of differing lengths, allowing the sequence to be read in a chromatogram

(Sanger et al., 1977). A disadvantage of using Sanger sequencing is that it can only run one sequence at a time. On the other hand, as an advantage, Sanger sequencing can run up to 1,000 base pairs at a time. Because of these attributes this sequencing method has been the predominant method for sequencing and characterising bacterial communities using 16S gene clone libraries (Tewari et al., 2011). However, a high-throughput technology called Next

Generation Sequencing (NGS) has recently been made available for sequencing more than one sequence in parallel (Anderson and Schrijver, 2010). This breakthrough supersedes Sanger sequencing in whole microbiome investigations.

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1.4.3 Next Generation Sequencing (NGS).

NGS is a technology which enables the sequencing of multiple small fragments of DNA or

RNA in parallel. NGS has two main types: shotgun sequencing and amplicon sequencing.

The type of nucleic acid that is sequenced and the read lengths, are dependent upon the platform used. Shotgun sequencing is used for whole genome sequencing, while amplicon sequencing targets a specific gene type and adds unique indexes into amplicons, allowing multiple samples to be sequenced simultaneously. Shotgun sequencing is substantially more expensive however, as an advantage, it does not rely upon PCR, and thus sidesteps a possible pitfall since PCR can fail to be sensitive enough to amplify low copy number bacterial DNA

(Greay et al., 2018). Amplicon sequencing allows for the profiling of a whole bacterial microbiome without the need for a-priori hypothesis, unlike PCR and Sanger sequencing, and is less expensive and requires less coverage compared with shotgun sequencing.

Amplicon sequencing of a tick’s bacterial microbiome uses PCR, firstly, to amplify the bacterial 16S rRNA (16S) gene. The ubiquitous 16S gene has nine hypervariable regions

(V1-V9) flanked by conserved regions (Greay et al., 2018). The V1-V2 and V3-V5 are the most commonly used hypervariable regions for microbiome analyses, and the use of the middle section (V3-V5) allows for easy alignment with previously sequenced bacteria to determine phylogenies(Greay et al., 2018; Sperling et al., 2017; Zolnik et al., 2016).

However, one limitation of NGS is the short sequences which are generated. This results in poor taxonomic resolution to the species level in most taxa and thus, careful interpretation of genetic data is paramount in the discovery of new species or strains of tick-borne pathogens

(Dantas-Torres et al., 2013). To overcome this limitation, a secondary PCR is often employed to increase sequence length and provide improved phylogenies (Gofton et al.,

2015b; Loh et al., 2017).

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1.4.4 TBD In Australia

In Australia there are few known bacterial TBD in humans; the most commonly encountered include Queensland tick caused by Rickettsia australis, Flinders Island Spotted Fever caused by R. honei, and caused by Coxiella burnetii (Graves and Stenos, 2017). Up until the time of this review, Q fever is the only locally-acquired notifiable TBD in Australia

(Graves and Stenos, 2017). However, Q fever is not traditionally associated with a tick bite, although ticks are required to complete its sylvatic lifecycle. The Australian paralysis tick,

Ixodes holocyclus, is endemic on the east coast of Australia and has been associated with the transmission of the bacterial agents that cause Q fever and (Graves and Stenos, 2017). The ornate kangaroo tick, A. triguttatum, is found throughout most of northern, central and western Australia and has been associated with the transmission of C. burnetii and a proposed pathogenic spotted fever group Rickettsia; R. gravesii (Owen et al.,

2006), while the southern reptile tick, B. hydrosauri, which occurs mainly in south eastern

Australia, transmits R. honei, the causative agent of Flinders Island Spotted Fever (Stenos et al., 2003b).

1.4.5 Rickettsia

The genus Rickettsia (phylum Proteobacteria, class alpha-Proteobacteria; order Rickettsiales; family ) is comprised of small, Gram-negative, obligate intracellular, pleomorphic coccobacilli bacteria. Species of Rickettsia are frequently transmitted by ticks

(Portillo et al., 2017). Rickettsia has a highly conserved 16S gene region and is unable to be discriminated into a species level by this gene (Portillo et al., 2017). Because of this, the gltA, ompA and ompB genes are frequently used as targets for species identification in human and vector samples(Vilcins, 2009; Whiley et al., 2016). The genus can also be divided into species groups according to their genetic and antigenic characteristics, and the groups of this genus are as follows: the spotted fever group (SFG), the typhus group (TG) and the ancestral group,

14

which contains a single species, Rickettsia bellii (Stothard et al., 1994). Several of the TG and

SFG have been identified as causing disease in humans to varying degrees of morbidity

(Graves and Stenos, 2017).

The SFG group R. honei, the etiological agent of Flinders Island Spotted Fever, was the first

Rickettsia associated with a reptile tick, B. hydrosauri, after the tick was found parasitising humans (Stenos et al., 2003b). Rickettsiae have since been identified in reptile ticks as probable primary endosymbionts (Vilcins, 2009; Whiley et al., 2016). While B. hydrosauri is a generalist reptile tick, there are still a number of unknown answers regarding the true identity of the R. honei vector and whether bobtail lizards (T. rugosa) act as a reservoir for R. honei.

This is because B. hydrosauri rarely bite humans and furthermore PCR amplification of

Rickettsia from tick eggs indicate that this Rickettsia is maintained via vertical transmission

(Stenos et al., 2003). Thus, lizards are not thought to be the reservoir, although they are the main vertebrate host of B. hydrosauri (Stenos et al., 2005). Thus, more research is necessary to determine if the lizard may be a transient host as their metabolism differs from mammals, or if the bacterium is quiescent in tissues or organs rather than in blood (Whiley et al., 2016).

However, this is out of the scope of the current project.

1.4.6 Coxiella

The genus Coxiella (Phylum Proteobacteria, class gamma-Proteobacteria; order Legionellales; family Coxiellaceae), are small, Gram-negative bacteria(Angelakis and Raoult, 2010).Based on the 16S gene, the bacterium was reclassified from the order Rickettsiales to Legionellales, within the gamma proteobacteria group, and the bacterium’s sister taxa include Legionella spp., , and Rickettsiella spp. (Raoult et al., 2005).

Coxiella burnetii causes Q fever in humans, which is a worldwide zoonosis with the exception of New Zealand and Antarctica, while in animals it causes coxielliosis (Duron et al., 2015).

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Animals infected with C. burnetii can act as carriers and shed it in urine, faeces, colostrum and milk intermittently over long periods. The human disease Q fever is typically acquired through the inhalation of contaminated aerosols the amniotic fluid or placenta, or the contaminated wool of livestock, which are the main reservoirs, or from soil (Angelakis and

Raoult, 2010). Coxielliosis in animals is generally an asymptomatic to mild or subclinical clinical disease; however, abortions are also known to occur. Furthermore, a wide variety of native marsupials and rodents show evidence of infection (Banazis et al., 2010; Bennett et al.,

2011; Cooper et al., 2012; Glazebrook et al., 1978).

Phylogenetic analysis of the Coxiella genus show an extensive genetic diversity within at least four divergent clades (Duron et al., 2015). Coxiella like endosymbionts (CLE) are found in all the clades however C. burnetti form a low genetically diverse cluster, indicating a unique and recent emergence of this pathogen (Bonnet et al., 2017; Oskam et al., 2017). Due to the phylogenetic pattern of the genus, C. burnetti is considered to have evolved from a CLE ancestor which was able to infect veterbrate cells (Duron et al., 2015).

1.4.7 Francisella

The genus Francisella (phylum Proteobacteria, class ; order

Thiotrichales; family Francisellaceae) consists of short, coccobacilli bacteria, which are frequently transmitted by ticks (Challacombe et al., 2017).The bacterium Francisella tularensis is the causative agent of Tularaemia, which is a highly infectious zoonotic disease

(Foley and Nieto, 2010). Tularaemia can be acquired by the bite, scratch or ingestion of a diseased animal, ingestion of or contact with infected water, aerosol inhalation and by ticks

(Foley and Nieto, 2010). While tularaemia is a notifiable disease, it is considered exotic in

Australia and there are no reports of an autochthonous infection (Eden et al., n.d.). However, there was recently a documented case of bacteraemia caused by a Francisella-like species in

Western Australia of unknown aetiology (Aravena-Román et al., 2015). The species genotypically resembled F. hispaniensis, which was isolated from patients in Spain (Huber et

16

al., 2010). There have also been Francisella-like endosymbionts (FLEs) sequenced from ticks parasitising reptiles in Thailand and the Northern Territory of Australia (Sumrandee et al.,

2014; Vilcins, 2009). These FLEs were revealed to be closely related to the pathogenic forms in phylogenetic trees.

1.5 MICROBES ASSOCIATED WITH REPTILES

Salmonella enterica and are the only pathogenic bacteria that have been positively identified from bobtails at the time of this review (Norval et al., 2019). These enteric bacteria are thought to be normal intestinal flora for the lizards, although they can also cause disease in humans (Whiley et al., 2017). Another group, the Mycobacteria, have several species which have been isolated after causing disease and fatality in reptiles, from the environment, and living as commensals in healthy reptiles (Mitchell, 2012). These species include M. chelonae, M. avium, M. schlangen, M. tropidonotus, M. marinum, M. thamnopheos and M. xenopi. Lizard species have been implicated as reservoirs for M. ulcerans, a disease of humans in Australia and Africa (Mitchell, 2012). Mycobacterium are thought to gain access to the reptile via cutaneous routes then spread systemically. Importantly ticks have recently been implicated as having a possible role in the transmission of Mycobacterium in the

Northern hemisphere (Palomar et al., 2019).

1.5.1 Zoonotic disease

Research into potential zoonotic diseases is important as the human encroachment on the natural habitats of wildlife increases (Dantas-Torres et al., 2013; Spratt, 2005). This increase of human wildlife interaction also increases potential for vector-borne diseases. In the case of mosquitoes, another important hematophagous vector, there was evidence that proximity to the vector in the peel region of Western Australia increased the chance of Ross

River virus (Jardine et al., 2015). Additionally, because of the continued development in the

Perth area, human dwellings could also encroach on areas where ticks are endemic and

17

uncharacterised as biting humans or where the vertebrate reservoir is potentially amplifying infectious agents (Dantas-Torres et al., 2013). Therefore, this warrants further research into the endemic ticks of Australia.

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1.5.2 Hypotheses and Aims

Ticks which parasitise T. rugosa are frequently surveyed in South Australia, however there is less knowledge about those infesting the lizard in Western Australia. This current research will expand upon the morphological and molecular taxonomy of bobtail ticks focusing on those found in Western Australia. This will be carried out by utilising ticks collected by wildlife health centres over the course of approximately seven years. There is a lack of knowledge about the bacterial communities in wildlife ticks in Australia and reptile ticks have been implicated as a human disease vector. Studies into parasites which are associated with wildlife can also be used for proactive measures in managing existing and emerging diseases. This study will be the first to sequence the entire bacterial microbiome in

Australian lizard tick.

The first hypothesis is that the ticks will be from one or more of the three commonly associated bobtail ticks, A.limbatum, A. albolimbatum and B. hydrosauri. The second hypothesis is that the immature life stages will be able to be molecularly identified using the CO1 gene. The third hypothesis of this project is bacterial communities within different tick instars will be unique and that there will be novel bacterial species within the ticks.

This present study aims to i) morphologically and molecularly identify ticks from

Australian reptiles focusing on Western Australian bobtails, Tiliqua rugosa rugosa; ii) characterise the bacterial diversity using the NGS Illumina Miseq platform; and iii) conduct a phylogenetic analysis on taxa of interest.

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Chapter 2: Materials and Methods

2.1 Sample collection & identification

Reptile ticks were collected in Western Australia by members of the public, Armadale reptile centre, GeoVet Busselton and Kanyana Wildlife Rehabilitation Centre. The ticks were all collected opportunistically from bobtails and were submitted to the Vector and Waterborne

Pathogen Research Group at Murdoch University. The majority of the ticks were stated to be from within the Perth metropolitan area. The ticks were stored in 70% ethanol tubes according to their individual host and stored at 4°C until ready for identification.

In preparation for identification, forceps and probes were sterilised in a sterile cabinet SC201

UV light box, then prior to individual tick identifications, probes and forceps were immersed in a 10% bleach solution and DNA AWAY TM (Molecular Bioproducts Inc., San Diego, USA) solution to decontaminate. Ticks were removed from 70% ethanol and examined in petri dishes and observed under an Eclipse E200 compound microscope (Nikon Corporation, Tokyo,

Japan) and an Olympus SZ61 stereo microscope (Olympus, Centre Valley, PA, USA).

Identification keys for Australian ticks (Barker and Walker, 2014; Keirans et al., 1996, 1994;

Roberts, 1969; Roberts, 1970) were used to determine species, instar and sex. Tick feeding status was also recorded; ticks were considered engorged if their conscutum was a bulbous, oval shape.

Following morphological identification, ticks of the same host registration were stored according to instar in new collection tubes containing 70% ethanol and at 4°C awaiting molecular analysis. In most cases nymphs and larvae could not be accurately identified to species level, in these cases samples were assigned to genus level. For these specimen’s molecular identification was attempted, see section 2.3. Analysis getIT photography software

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(Olympus, Centre Valley, PA, USA) was utilised to photograph specimens of interest and illustrations were made of defining features.

2.2 DNA extraction

Prior to DNA extraction, individual ticks were surface-sterilised with 10% sodium hypochlorite, then 70% ethanol, and washed with sterile DNA-free Phosphate-Buffered saline

(PBS) and air-dried. DNA-free equipment and tubes were used for each step and equipment was decontaminated between samples with DNA AWAY TM (Molecular Bio-products Inc.,

San Diego, USA). Genomic DNA (gDNA) was extracted using the DNeasy Blood and Tissue

Kit (Qiagen, Germany) following the manufacturers’ recommendations (Qiagen

Supplementary Protocol: Purification of total DNA from insects), with modifications.

Following the surface decontamination adults were bisected and half was placed back into ethanol tube for future analysis; in the case of immature life stages the whole specimen was used. Ticks were placed in a 2 mL safe lock tube (Eppendorf TM) with a 5 mm steel bead and immersed in liquid nitrogen for approximately 5 mins. After snap freezing the samples were homogenised using a Tissue Lyser LT (Qiagen, Germany) for 1-3 mins at 40 Hz. The buffer

ATL and Proteinase K (pk) enzyme were then added to the tubes at 450 μL buffer and 50 μL pk for engorged adults, and 200 μL and 22.5 μL pk for nymphs, larvae and unengorged adults.

Samples were then incubated on a heat block at 56°C, 700 rpm, for 16 to 18 hours. Following incubation 200 μL of supernatant was transferred into a sterile 1.5 mL safe lock tube

(Eppendorf TM) with 200 μL of 96% ethanol and 200 μL of buffer AL and then vortexed thoroughly. Following 500 μL buffer AW1 and buffer AW2 washes buffer AE was added directly to the membrane in the following volumes: engorged adults 150 μL, unengorged adults 60-75 μL and larvae and nymphs 40 μL. A double elution was utilised for all unengorged adults, nymph and larvae. Samples were incubated at room temperature for 4 minutes and then centrifuged at 8,000 rpm for 1 min.

22

Extracted gDNA was stored at -20°C until molecular analysis. Twenty of the samples were extracted without an extraction blank, for all further DNA extractions an extraction reagent blanks (EXB) (n=3) were performed in parallel with the DNA extractions. This was in order to establish background bacterial populations. Extraction steps were undertaken wearing gowns, gloves, plastic sleeves and face masks to minimise contamination by humans and the environment.

2.3 Amplification of the Ixodida cytochrome oxidase subunit 1 (CO1) gene

Two female Amblyomma albolimbatum ticks were used as molecular reference specimens to generate genetic data. The reference specimens were chosen based on morphological identification (see results for photograph/composite illustration) and geographic position, northern reference specimen R76 (-31.756999, 115.813270) and southern reference specimen

R1 (-33.654972, 115.330620). A subset of morphologically divergent adults, nymphs and larvae (n=7) was sequenced to confirm taxonomic identity of ticks.

Table 2.1. Primers targeting the cytochrome oxidase subunit 1 (CO1) gene of ticks

(Ixodida).

Primer Sequence (5` -3`) Reference

HCO2064 GGT GGG CTC ATA CAA TAA ATC C Song et al., 2011

HCO1240 CCA CAA ATC ATA AAG ACA TTG G

The molecular barcoding gene CO1 was chosen based on previous research by Evans et al

(2019), which identified this as a useful gene for species delimitation of Australian Ixodida.

The PCR assay was carried out according to Evans et al (2019) utilising primers from Song et al (2011) (table 2.1) to amplify an ~850bp fragment of the CO1 gene. Reactions were carried out in 25 μl volumes consisting of 1X KAPA buffer with dye (KAPA Biosystems,

Massachussetts, USA) 2.0 mM MgCl2, 0.5 mM of each dNTP (FisherBiotech, Australia), 0.12

23

μM of each primer (table 2.1), 1.25 U KAPA Taq (KAPA Biosystems, Massachussetts, USA) and 1 μl of neat gDNA. Thermal cycling was performed on a BioRAD T100 under the following conditions; an initial denaturation of 95°C for 2 mins, followed by 35 cycles of 95°C for 30 s, 48°C for 30 s and 72°C for 50 s with a final extension of 72°C for 2 mins. No template controls consisting of molecular grade water were included in each assay. Amplicon products were visualised on a 1% agarose gel, and bands of the correct size were excised and sequenced using the forward primer (HCO2064) as outlined in section 2.6.

2.3.2 Phylogeny

Sequences were imported into Geneious v10 (Kearse et al., 2012) for inspection and trimming.

The sequences were then aligned with a selection of Australian Ixodidae ticks reference sequences, retrieved from GenBank (Benson et al., 2017), using with the parameters set out in

Evans et al (2019). Aligned sequences were then imported into MEGA7 (Kumar et al., 2016) and the most appropriate nucleotide substitution model was chosen based on the lowest

Bayesian Information Criterion (BIC) score.

2.4 Library preparation and NGS

Next-generation sequencing (NGS) was carried out on a sub-sample of bobtail ticks (n=116), on the Illumina MiSeq platform, targeting a ~460 bp product of the V3-4 hyper-variable region of the bacterial 16S gene (Table 2.2). Extraction reagent blanks (EXB) (n=3) and no-template controls (NTC) (n=3) were included at all stages to quantify environmental microbial communities. Library preparation and sequencing was carried out following Illumina 16S metagenomic sequencing protocol (#15044223 Rev B) with some modification. All pre-PCR and post-PCR procedures were performed in physically separate dedicated laboratories, and strict hygiene protocols were maintained through library preparation in order to minimise amplicon contamination.

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2.4.1 Amplicon PCR

The first amplicon stage of the library preparation was carried out using the primers set out in table 2.2 in 25μl reactions. Each reaction contained the following: 1X KAPA buffer with dye

(KAPA Biosystems, Massachussetts, USA) 2.0 mM KAPA MgCl2, 0.4 mg/ml BSA

(FisherBiotech, Australia), 0.25 mM of each dNTP (FisherBiotech, Australia), 0.4 uM of each primer containing Illuma MiSeq adapters (table 2.2), 0.5 U KAPA Taq (KAPA Biosystems,

Massachussetts, USA) and 2 μl of neat gDNA. Controls (EXBs and NTCs) were included in each assay. Thermal cycling was performed on a BioRAD T100 under the following conditions: initial denaturation at 95°C for 3 mins, followed by 35 cycles of 95°C for 30 s,

55°C for 30 s, 72°C for 30 s and a final extension of 72°C for 5 mins. Amplification was confirmed by visualising 5 μl of the PCR products on a 2% agarose gel, with a 100bp molecular weight ladder.

25

Table 2.2 Primers used for 16S gene bacterial next-generation sequencing amplicon assays.

Primer Sequence (5`-3`) Reference

338F ACT CCT ACG GGA GGC AGC AG (Lopez et al., 2003)

806R GGA CTA CHV GGG TWT CT AAT (Caporaso et al.,

2011)

MiSeq forward TCG TCG GCA GCG TCA GAT GTG TAT AAG (Illumina Inc, 2015)

adapter AGA CAG

MiSeq reverse GTC TCG TGG GCT CGG AGA TGT GTA TAA

adapter GAG ACAG

2.3.3 Index PCR and gel electrophoresis

Amplicons were indexed using the Nextera XT Index 1 primer (N7XX) and Index 2 primer

(S5XX). The assay was made up into 25 μl reaction volumes containing 12.5 μl of 2X KAPA

HiFi HotStart Ready mix, 2.5 μl of each index primer and 1μl of the amplicon PCR product.

Thermal cycling was performed on a BioRAD T100 under the following conditions: 95°C for

3 mins, 15 cycles of 95°C for 30 s, 55°C for 30 s, 75°C for 30 s and a final extension at 72°C for 5 mins. Following PCR 2 μl of 6X loading dye (ThermoFisher Scientific, California USA) was mixed with 5 μl of indexed PCR product and run on a 2% agarose gel, along with a 100 bp molecular weight ladder. Index PCR products were visualized to ensure the first-round product had increased by ~50 bp to confirm successful index binding.

2.3.4 Index PCR clean up and gel electrophoresis

Indexed samples were purified by following using Agencourt® AMPure® XP PCR

Purification beads (Massachusetts, USA) following manufacturers recommendations with minor modifications. In order to remove unwanted DNA products (below 300 bp) 10 μl of the

26

index PCR product was mixed with 8 μl of AMPure beads. Following 5 mins incubation at room temperature, the plate was exposed to a magnetic field for 2 mins. The supernatant was removed and followed by two washes with 80% ethanol. After all ethanol was aspirated an elution was performed using 30μl of QIAGEN EB buffer. Following purification 5 μl of each sample was mixed with 2 μL of 6 X loading dye (ThermoFisher Scientific, California USA) and separated on a 2% (w/v) agarose gel with a 100 bp molecular weight ladder (Promega,

Madison USA). The images from this gel electrophoresis were used to quantify samples in preparation for library pooling. The remainder 25 μl of the eluted DNA was stored at -20°C until further analysis.

2.3.5 Library pooling and quantification

Following gel electrophoresis of purified index products samples were pooled into five libraries based on band intensity. To ensure equimolar concentration of pooled samples the intensity of purified index products was calculated using GelAnalyzer (Lazar and Lazar,

2010). Libraries were then quantified using the Qubit® 2.0 Fluorometer (Thermo Fisher,

Australia) following the manufacturers’ protocol. A final library of 10 mM was then sent to the Australian Genome Research Facility, Melbourne for sequencing on the Illumina Miseq platform. A final library of 9 pM containing 15% PhiX control was sequenced using v3 chemistry on a 2x300 cycle paired end kit.

2.4.1 Bioinformatics

Raw fasta files were downloaded from the Illumina BaseSpace Sequence Hub for analysis in a USEARCH v11 (Edgar, 2010) pipeline. Raw paired reads were first merged and with a minimum merge overlap of 50 bp. Only sequences with perfect primer matches were retained and then the primers and distal bases were removed. Sequences were then quality filtered, allowing a <1% expected error rate and singletons were discarded (Edgar, 2016). Sequences were then clustered into zero radius-operational taxonomic units (ZOTUs) using the UNOISE3 algorithm (Edgar, 2016). Taxonomy was assigned in Quantitative Insights into Microbial

27

Ecology (QIIME2 v2019.4) using the q2-feature-classifier (Bokulich et al., 2018) with reference to a trained Greengenes database (DeSantis et al., 2006) (release May 2013).

Taxonomic assignments were confirmed using NCBI blast (Abdad et al., 2017)The profiles of EXB and NTC samples were first inspected to ensure quality of samples and then bioinformatically to eliminate potential contaminating background bacterial sequences.

2.4.2 Ecological Modelling

The bacterial diversity was assessed using QIIME (v2019.4). Alpha diversity analysis included rarefaction curves and plots using OTU richness, Shannon’s diversity metric and Faith’s phylogenetic diversity. Diversity metrics were separately compared among instars and seasons with Kruskal-Wallis tests. To assess the variability of OTU richness among instars, the communities were visualised using principal coordinate analysis emperor plots of beta diversity with the Jaccard index (presence/absence) and the Bray-Curtis dissimilarity index

(abundance). These diversity metrics were further used to compare the instars bacterial communities with permutational analyses of variance (PERMANOVA). PERMANOVAs were performed with 999 permutations among all instars and pairwise PERMANOVAs were performed between instars with 999 permutations, with Benjamini Hochberg adjustment of P- values. Further percentage abundance analyses were visualised using the Ampvis package in

Rstudio (v1.2.1335). The fed versus engorged status of females was compared for the top two most abundant ZOTUs using t-test, which was carried out in Excel.

2.5 Detection of Coxiella using target PCR

A conventional PCR assay was employed to screen samples (n=116) for Coxiella, targeting a

524bp fragment of the 16S gene (short 16S). This was performed using the primers Cox- sp434F/Cox-sp1004R(Lalzar et al., 2012). Any samples that produced a product visible on an

28

agarose gel, and samples that were positive for Coxiella sp. via 16S NGS bacterial profiling, were then subject to a second Coxiella genus-specific PCR targeting 1.45 kb of the 16S gene

(long 16S) using the primer pair Cox-16S-1457F/Cox-16s-1457R (Masuzawa et al., 1997).

Reactions were carried out in 25μl volumes each containing: 1X KAPA buffer with dye

(KAPA Biosystems, Massachussetts, USA) 2.5mM KAPA MgCl2, 0.25mM of each dNTP

(FisherBiotech, Australia), 0.4μM of each primer, 0.5 U KAPA Taq (KAPA Biosystems,

Massachussetts, USA) and 2μl of neat gDNA. A PCR no-template control (NTC) of 2μL of molecular grade water was used in each assay. Thermal cycling was performed on a BioRAD

T100 under the following conditions for the short 16S assay: initial denaturation at 95°C for 5 mins, followed by 40 cycles of 95°C for 30 s, 58°C for 30 s, 72°C for 30 s and a final extension of 72°C for 5 mins. Thermal cycling conditions for the long 16S assay: initial denaturation at

95°C for 5 mins, followed by 40 cycles of 95°C for 30 s, 48°C for 30 s, 72°C for 1 min and a final extension of 72°C for 5 mins. Amplicons were visualised on an agarose gel as outlined in section 2.6. To test for inhibition in PCR assays various DNA concentrations were tested either containing 4μl neat, 2μl neat, 2μl 1:10 diluted, 2μl 1:100 diluted.

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Table 2.3 Primers used for Coxiella genus specific PCR assays.

Primer Sequence (5` -3`) Amplicon Reference

(bp)

Short 16S

Cox-sp434F CCT TTT GAG CGT TGA CGT TA ~524 Lalzar et al., 2012

Cox-sp1004R CCA AAG GCA CCA AGT CAT T

Long 16S

Cox-16s- ATT GAA GAG TTT GAT TCT GG ~1450 Masuzawa et al.,

1457F 1997

Cox-16s- CGG CTT CCC GAA GGT TAG

1457R

2.6 Gel electrophoresis and Sanger sequencing

Nested and conventional PCR products were separated on a 1-2% agarose gel run in 1 X tris- borate-EDTA (TAE) buffer and stained with SYBR Safe (Invitrogen, Australia) for examination with UV transillumination. All products were visualised with a 100 bp or 1 kb molecular weight ladder (Promega, Madison USA) and bands of the desired size were excised using sterile scalpels. Products were purified for sequencing using an in-house filter tip method. A sequencing mixture containing a volume of 4-6μl of PCR product, Milli-Q water, and 1μl of corresponding forward primer totalling 12μl were made for each positive sample selected. Sanger sequencing was performed at the Australian Genomic Research Facility

(Perth, Australia) on an Applied Biosystems 3730 capillary sequencer using Big Dye

Terminator chemistry v3.1.

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2.7 Phylogenetics

Sequences were imported into Geneious v10 (Kearse et al., 2012)and primers were trimmed.

Sequences were aligned with reference sequences retrieved from GenBank using MAFFT v7.017(Katoh and Standley, 2013)and realigned using MUSCLE(Edgar, 2004). An outgroup was used to root trees and provide a more meaningful comparison due to the short fragment length of sequences analysed. Trees were constructed using the lowest BIC score.

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Chapter 3: Results

3.1 TAXONOMIC IDENTIFICATION OF TILIQUA RUGOSA TICKS

3.1.1 Morphological Identification

A total of 306 ticks from approximately 80 bobtail lizards (Tiliqua rugosa) in Western

Australia (fig. 3.1) were morphologically identified. The approximation of the number of hosts necessary in light of the collection methodology, where separate tubes of ticks were filled from the same host or single tubes from multiple hosts, and observer record keeping variations.

There were a number of ticks from an unknown locality in Western Australia and these were not included in further analysis (n=30). The most frequently identified instar was male at 43.1% (n=119), followed by nymphs 40.6% (n=112) and 15.2% (n=42) were females, while only three larval ticks (1.1%) were observed. Ticks from all stages of engorgement were recorded (fig. 3.2).

All ticks were morphologically identified as belonging to the genus Amblyomma. This was determined based on the following anatomical features: posterior anal groove, elongated palps, eyes present and large pale yellow to light brown orange in colour, ornate scutum and well-developed festoons (fig.3.3). Ten ticks from the Busselton area (n=25) were morphologically distinct. The ticks that were morphologically distinct had dark brown to black eyes, which were not obvious at first identification (fig. 3.4). Furthermore, these ticks also had a much darker background colour to the other ticks. However, all other morphological traits were consistent with being the genus Amblyomma. Several ticks also showed iridescent ornamentation on the scutum, which is morphological feature of Amblyomma ticks (fig 3.4).

Several morphological features were used to identify ticks to the species level.

Amblyomma albolimbatum was identified by the following morphological traits: mild

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expansion of the posterolateral angle of the scutum; 2 (3.3 dorsal view arrow 1); numerous uneven punctations on the scutum, many of which were course; the two porose areas on the basis capitulum were large, broadly oval, which were centrally spaced at an interval of approximately one and a half the diameter of a porose area (dorsal view arrow 2); eyes that were large, pale and flat (dorsal view arrow 3); dark bandings on the articulations of the leg segments (dorsal view arrow 4); sparse and scattered hairs; presence of two uneven spurs on coxa I with a longer external spur (ventral view arrow 1 ); a single spur on subsequent coxae; longer and more elongated spur on coxa IV than the external spur on coxa I; spiracular plate that was broadly comma shaped (ventral view arrow 3); genital aperture adjacent to coxa II

(ventral view arrow 4).

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Figure 3.1 Geographic distribution of ticks collected for this study. The large navy-blue circle shows the Northern representative specimens’ location (-31.756999, 115.813270) and the purple circle shows the Southern representative specimens’ location (-33.654972,

115.330620).

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Figure 3.2 Amblyomma nymphal ticks with varying degrees of engorgement (dorsal).

Nymphs were identified based on the absence of a ventral genital pore and the presence of four pairs of legs.

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Figure 3.3. Female Amblyomma albolimbatum (R76 Northern representative sample). Top)

Dorsal view 1. scutum; 2. porose areas on basis capitulum; 3. Eye; 4. Banding on legs.

Bottom) ventral view 1. Two spurs on coxa 1; 2. anal groove; 3. Spiracular plate; 4. Genital aperture 2. Left leg 1 was damaged postmortem.

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Figure 3.4 Male Amblyomma albolimbatum (PI1872) with iridescent ornamentation on the scutum; Dorsal view.

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Table 3.1 Instar of ticks and species morphologically identified from Western Australian

Tiliqua rugosa. Those designated for DNA extraction in brackets.

Female Male Nymph Larva

Amblyomma 36(26) 117(53) 95(13)

albolimbatum

Amblyomma spp. 6(3) (2) (17) 3(2)

DNA extraction 29 55 30 2

sum

The majority of the tick samples were collected in the late spring/summer months, November to January (70%) over the full range of years, and in 2014, ticks were collected in during the winter months, June to August. The tick samples selected for DNA extraction for ongoing molecular analyses spanned the following years: 2013 (12.9%), 2014 (41.4%), 2015 (26.7%),

2016 (8.6%), 2017 (0.9%), 2019 (9.5%). Ticks that were selected for DNA extraction included

55 males (47.4%), 29 females (25%), 30 nymphs (25.9%) and two larvae (1.7%) (tables 3.1 and 3.2; fig. 3.5).

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Figure 3.5 Amblyomma larval ticks (dorsal view. Left, pale unfed; Right, fed). Larvae were identified by the presence of three pairs of legs.

Table 3.2 The instar and status of the ticks that were morphologically identified and the number of those ticks which were chosen for for molecular analyses.

Number of Ticks Number of Ticks chosen Instar & Status Identified for DNA extraction Larva 3 2 Fed 1 1 Pale 2 1 Nymph 122 30 Engorged 50 26 Fed 54 3 Pale 18 1 Male 124 55 Fed 116 51 Pale 8 4 Female 57 29 Engorged 17 7 Fed 39 21 Pale 1 1 Total 306 116

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3.1.2 Molecular identification

Prior to 16S bacterial community profiling, two larvae and 17 nymphs of unknown taxa

(i.e. were removed from a host without male or female ticks present), were subjected to molecular barcoding PCRs to provide accurate species identification. In addition, two representative tick samples of known taxonomy, based on morphological identification in figs.

3.1 and 3.3, one from Southern WA and one from Northern WA, along with a subsample (n =

4) of the morphologically distinct samples from the Busselton area and additional seven A. albolimbatum specimens were included for a higher resolution, and a larger geographical range.

All 32 ticks, used for molecular identification, had their CO1 gene successfully amplified, showing the correct band size upon gel electrophoresis. However, only nine (28%) were successfully sequenced. A sequence was considered successful if it had a clean chromatogram consisting of characteristic single peaks for each nucleotide. Only 1/17 nymph and 2/2 larvae were successfully sequenced; of the reference samples, 4/12 female and 2/2 male ticks were sequenced. The limited success of the sequencing was due to laboratory contamination rather than samples of mixed origin, however due to time and budget constraints, the samples were not re-amplified.

A single phylogenetic tree (fig 3. 6) was constructed as a result of the limited number of reference sequences available on GenBank. This tree comprised a 425bp alignment with sister taxa, A. fimbriatum, A. triguttatum and A. limbatum (fig 3.6). This tree shows high bootstrap support for the A. albolimbatum monophyletic clade, with the Northern and Southern representative sequences split across two branches. The two larvae clustered with the southern branch and the nymphs clustered with the Northern reference sample. The four morphologically distinct adult sample sequences clustered with the Southern reference specimens with no genetic difference.

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Figure 3.6 shows a monophyletic clade produced by the Amblyomma albolimbatum samples sequenced in this study. There are two branches which coincide with the Northern and Southern representative samples. The GTR model percentage intraspecific distance compared in table 3.3 shows a high relative pairwise distance between the Northern and

Southern representative samples (R76 & R1) of 4.79%. The highest pairwise distance is 5.06% between the Northern representative sample R76 and R120. The sample R120 also diverges from the Southern branch slightly at 0.24% and R118 diverges from the Northern branch at

0.48%.

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Figure 3.6 Maximum likelihood phylogenetic tree based on 425bp CO1 alignment of two reference specimens of Amblyomma albolimbatum species identified within this study against specimens of unknown morphology. The phylogenetic tree was built using the Hasegawa-

Kishino-Yano model. The trees were rooted with soft ticks of the genus Argas. The species are compared to other Australian hard ticks (GenBank accession numbers shown). Branch lengths indicate the number of character changes and all bootstrap values from 1,000 replications are shown on interior branch nodes.

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Table 3.3 The percentage pairwise distance between all samples sequenced that make up the

monophyletic Amblyomma albolimbatum clade. The Reference samples are denoted by

“North”, “South”.

R76F R118N R102L R120F R101L R87M R86F R45M North

R76F North 0.48

R102L 4.78 4.79

R120F 5.05 5.06 0.24

R101L 4.78 4.79 0 0.24

R87M 4.78 4.79 0 0.24 0

R86F 4.78 4.79 0 0.24 0 0

R45M 4.78 4.79 0 0.24 0 0 0

R1F South 4.78 4.79 0 0.24 0 0 0 0

The ticks formed a monophyletic clade apical to the other Amblyomma species with

COI data. The were two branches in the clade which were made up primarily of the Northern

and Southern samples. However, the larvae, which were from the North ,clustered with the

Southern samples. There was a 4.79% pairwiase distance between the two representative

samples. One tick R120 had a genetic divergence form R76 of 5.05% and from R1 of 0.24%.

The genetic distance between the A. albolimbatum sequences from this study were highest

with A. fimbratum (table 3.4) and lowest with the morphologically similar species A. limbatum.

Although the other morphologically similar species A. vikkiri is reported to have the CO1 gene

on the NCBI database, the available sequence had a short 49bp overlap with the sequences

generated in this study.

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Table 3.4 The percentage interspecific differences, with standard error, between the

Australian Amblyomma species cytochrome oxidase genes available on NCBI database and those Amblyomma albolimbatum samples sequenced in this study

All (N+S) Northern Southern

A. albolimbatum A. albolimbatum A. albolimbatum

A. limbatum 9.14 ± 0.04 9.53 ± 0.04 8.98 ± 0.03

A. triguttatum 14.93 ±0.24 17.92 ± 2.50 14.57 ± 0.05

A. fimbriatum 23.70 ± 0.16 21.33 ± 2.43 23.92 ± 2.43

3.2 NEXT GENERATION SEQUENCING DATA EXPLORATION

NGS was carried out on the 116 ticks, which were chosen to represent different life stages, geographic distribution and year collected available in the collection and included six negative controls (three EXB and three NTC controls). The number of reads that passed filter set at Q30, indicating a 0.1 % chance of an incorrect base being called during sequencing, was a total of 9,352,906 raw pair-end reads were obtained for tick samples (n=116) and 354,014 reads from negative controls (EXB and NTCs) (n=6). After merging forward and reverse reads, 6,878,700 (97.3%) and 194,324 (2.7%) reads were retained for tick samples and controls, respectively. 3,694,305 and 65,423 reads were subsequently obtained upon quality filtering and trimming of sequences from tick samples and controls, respectively. Lastly, reads were clustered and 1,438 ZOTUs were generated.

ZOTUs observed in negative controls were considered ‘environmental’ contamination and co-amplified chloroplast and mitochondrial assigned ZOTUs were filtered out of the tick data set returning 3,658,365 reads. This resulted in a final total of 1,385 ZOTUs in ticks. In addition, when the sequencing depth threshold was increased to >30 reads per ZOTU, the total

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number of ZOTUs was reduced to 860, and at > 500 reads the total number of ZOTUs was reduced to 200.

3.2.1 Sequencing depth

The distribution of sequencing depth was not evenly distributed with the most reads in a single sample recorded as 96,464 from an unengorged female (R68) and the least recorded as 49 reads from an engorged nymph (R114). In addition to R114, four other samples (R80,

R54, R81, and R117) had less than 1,000 sequences, R114, R80, R54 and R81 containing less than 500 reads which is considered extremely low sequencing depth for this sample type.

The depth of sequencing and the species diversity followed a broad trend whereby increased depth correlated with increased diversity (Appendix table 2). However, the highest species diversity came from an unengorged female (R47) sample with only 34,018 reads and

310 ZOTUs, although when the sequencing depth threshold was increased (>30 reads per

ZOTU) the total number of ZOTUs dropped to 63. A male sample (R71) with a high number of reads (89,144) had an extremely high species diversity with 303 ZOTUs; when the threshold of number of reads per ZOTU was increased to >30, 130 ZOTUs remained. Additionally, the samples that had below 500 reads had seven ZOTUs. The average number of reads from the tick samples (n=116) was 31,537.63 ± 1773.56 (SD=19,101.82). The control samples (n=6) had an average number of reads equalling 10,903.83 ± 7047.49218 (SD=17262.76). The average number of ZOTUs for the samples was 53.63 ± 5.19 (SD= 51.44) and for the controls

19.17 ± 5.19 (SD=12.70).

The instars had a varied sequencing depth and only females and nymphs achieved a plateau of approximately 15,000 reads (Fig 3.7). Samples were rarefied to 15,000 reads due to the high variability between instars and in an effort to retain as many samples as possible. This rarefication retained 39.10% of the diversity within 80.33% of the samples.

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Figure 3.7 The rarefication curves plotted by the faith phylogenetic diversity metric, separated by instar, females (dark blue), nymphs (light orange) and control (green) show an adequate sequencing depth by displaying the characteristic plateau. Males (dark orange) and larvae

(light blue) do not plateau.

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3.2.2 Alpha and beta diversity among instars

All alpha diversity metrics (Fig. 3.8) showed significant differences between instar bacterial diversity (Kruskal Wallis, P<0.05). Significant differences were observed for pairwise test metrics between females and males; Faiths phylogenetic diversity metric (P= 0.008), Observed

OTUs (P<0.0016) and Shannon’s metrics (P<0.0000018). Females and nymphs also showed a significant difference for the Faiths phylogenetic diversity metric (p= 0.01). All other metrics did not report any statistically significant pairwise results.

The effect of presence/absence and abundance data was analysed through diversity models and plotted through principle coordinate analysis (PCoA) (Fig. 3.9). Overall, the Jaccard (J) and Bray Curtis (BC) pairwise PERMANOVA analyses provided consistent insight into the beta diversity between instars revealing significant differences between the bacterial compositions of females and males (J P=0.004; BC P=0.002), females and nymphs (J P=0.004;

BC P=0.002), males and nymphs (J P=0.012; BC P=0.002) and larvae and males (J P=0.004;

BC P=0.0495).

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Figure 3.8: Alpha diversity plots. Upper: Faiths phylogenetic diversity metric X2=11.39, P=

0.00975; Middle: Observed OTUs X2=16.94, P= 0.00073; Lower: Shannon’s X2= 28.8786,

P= 0.0000024.

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Female

Larva

Male

Nymph

Figure 3.9 PCoA analysis showing the differences in beta-diversity between tick instars at

ZOTU taxa between presence/absence data based on Jaccard index (left) and abundance data based on Bray-Curtis dissimilarity (right)

3.2.3 Microbial Composition

The bacterial microbiome of bobtail tick samples was dominated by the phylum Proteobacteria

(90.3%) as were the control samples (79.1%) (Fig 3.10). The next most dominant phyla in the tick samples was Actinobacteria (4.1%) and Firmicutes (3.4%) and there was also an abundance of over 1% of unassigned phyla, all other phyla were less than 1%. The negative control samples had a much higher abundance of Firmicutes (10.4%) and Actinobacteria (8%) sequences. Cyanobacteria (1.3%) was also observed.

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Figure 3.10 The abundance of phyla in the Amblyomma albolimbatum ticks samples and the

control samples. Abundance was calculated by number of reads in each ZOTU.

Francisellaceae 64.1 88.3 44.4 34.5

Rickettsiaceae 25.4 5.7 35.2 41.5

Coxiellaceae 0.2 0 3.3 2.9

Enterobacteriaceae 2 0.1 2 0.7

Planococcaceae 0.5 0.1 2.1 1.6

Bacillaceae 0.6 0 2.5 0.2

Brevibacteriaceae 0.4 0 1.3 0

p__Unnasigned_Otu212 2.8 0 0 0

Sphingomonadaceae 0.1 0 0.1 2.3

Moraxellaceae 0.2 0.1 0.9 0.2

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N F Figure 3.11. Family level taxonomic assignment for each instar of the tick Amblyomma

albolimbatum.

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Taxonomic assignment at the Order level were represented largely dominated by and Rickettsiales. Analysis at the Family level further divided tick instars by their unique microbiomes (fig 3.11). At the Family level, Francisellaceae were highly abundant in larvae

88.3% and females 64.1%, and, to a lesser extent, in males 44.4%, and nymphs 34.5%.

Rickettsiaceae were the most abundant family in nymphs 41.5% and the second most abundant in all other instars. Coxiellaceae, Sphingomonadaceae, , Planococcaceae and Bacillaceae were the other dominant Family taxa present in the ticks. ZOTUs sequences were edited manually to display the correct genus; for example, 24 ZOTUs were initially assigned to the order Legionellales instead of Thiotrichales by the GreenGenes database.

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Francisella 64.1 88.3 44.4 34.5 Rickettsia 25.4 5.7 35.2 41.5 Staphylococcus 1 0.1 2.3 1.5 Rickettsiella 0.2 0 2.4 2.1 Bacillus 0 0 2.2 0.1 Brevibacterium 0.6 0 1.8 0.2 Uncultured bacterium 0.7 0.2 0.7 2.1 Unnasigned 2.8 0 0.5 0.2 Coxiella 0 0 0.9 0.9 Pantoea 1.9 0 0 0.6 Enterobacter 0 0 1.1 0 Serratia 0 0 0.9 0 Kocuria 0.1 0.2 0.6 0.1 Sphingomonas 0.1 0 0.1 1.1 Psychrobacter 0 0.1 0.6 0 Paracoccus 0 0 0.2 0.7 c__Alphaproteobacteria_Otu1007 0.3 0.1 0.3 0.2 f__Sphingomonadaceae_Otu286 0 0 0 1.1 Halomonas 0 0 0 0.9 Pediococcus 0 0 0 0.9 Pseudomonas 0 0.1 0.1 0.5 Clostridium 0.3 2.5 0 0.2 c__Gammaproteobacteria_Otu482 0.2 0 0.2 0.1 Enterococcus 0.3 0 0.1 0.2 Roseomonas 0 0 0.3 0 c__Alphaproteobacteria_Otu279 0 0 0.3 0.1 Arthrobacter 0.1 0 0.3 0 Leucobacter 0 0.1 0.3 0.1 Acinetobacter 0.1 0 0.2 0.1 Sphingobacterium 0 0 0.2 0 Yushania 0 0 0 0.5 Flavobacterium 0 0 0 0.5 Dietzia 0 0 0.2 0 Streptococcus 0 0.1 0 0.4 f__Rhodobacteraceae_Otu674 0 0 0 0.5 Rhodococcus 0 0 0.2 0.1 Methylobacterium 0 0.1 0 0.4 Xanthomonas 0 0 0.2 0 p__Cyanobacteria_Otu37 0 0 0 0.4 Leifsonia 0 0 0 0.4 Patulibacter 0 0.1 0 0.4 Ruminococcus 0 0 0 0.3 Gordonia 0 0 0.2 0 Curtobacterium 0 0 0 0.3 f__Comamonadaceae_Otu894 0 0 0 0.3 Brachybacterium 0 0 0.1 0 Aureimonas 0 0 0 0.2 Corynebacterium 0 0 0.1 0 Pedobacter 0 0 0.1 0

o__Sphingomonadales_Otu383 0.2 0 0 0

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tick Amblyomma albolimbatum.

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Analysis at the Genus level showed the males and nymphs to have a more heterogenous environment than the females and larvae (fig 3.12). Francisella and Rickettsia were the most abundant at all life stages. The top two genera abundances were further visualised using boxplots (fig 3.13). Francisella were highly abundant in larvae 88.3% and females 64.1%, and to a lesser extent, in males 44.4%, and nymphs 34.5%. Rickettsia was the most abundant in nymphs 41.5% and second most abundant in all other instars. The two larvae exhibited a different relative abundance of Rickettsia; the pale only showed 0.6%, while the fed larvae presented 10.5 %. A number of other previously observed tick microbiota including Kocuria,

Pseudomonas, Brevibaterium, Pantoea, and Sphingomonas were identified at low relative abundance (<1%).

Francisella

Group

Nymph Male Larva Female

Rickettsia

0 25 50 75 100 Read Abundance (%)

Figure 3.12 Boxplots denoting the percentage relative abundance of the top two genera

Francisella and Rickettsia in each instar of the tick Amblyomma albolibatum.

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3.2.4 Microbial taxa of interest

NGS identified a number of ZOTUs assigned to bacteria known to be associated with ticks.

ZOTUs associated with taxa of interest were extracted from the dataset and individually compared against the NCBI database using BLAST (Table 3.6). As observed in Table 3.6, very few taxa of interest sequences were also observed in negative controls and likely represents cross-contamination of tick samples to controls. The percentage of cross- contamination was 0–0.023% within controls compared to total number of reads in the taxa of interest.

Several ZOTUs identified within A. albolimbatum ticks were genetically similar to previously described arthropod endosymbionts and tick-borne pathogens. The most abundant ZOTU

(ZOTU1) within A. albolimbatum ticks had a 100% sequence identity to a Rickettsia endosymbiont sequence (MK005802.1) from a tobacco whitefly, Bemisia tabaci. This

Rickettsia ZOTU was observed within 79.31% of ticks from all instars. ZOTU10 and ZOTU11 had high genetic similarity, >99.26%, to Rickettsia raoultii (MK304546.1). Rickettsia raoultii is a member of the spotted fever group and to date, has only been identified in Dermacentor ticks in the Northern hemisphere. Phylogenetic analysis was undertaken (fig 3.13) and all the

ZOTUs (1, 10 and 11) formed different branches, however no further conclusions can be made due to the small bp length.

The next most abundant ZOTU (ZOTU2) within A. albolimbatum ticks had a 99% sequence identity to a Francisella hispaniensis (KT281843.1) strain, which was recently reported in a human in WA with an unknown tick exposure (Aravena-Román et al 2015). This ZOTU had

100% prevalence in females and larvae and >90% in males and nymphs. However, the relative abundances differed with larvae showing 85.94% and 64.39% in females whereas nymphs showed 22% and 6% in males. No statistical difference was found in this ZOTU for fed vs

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engorged females (P=0.15). This F. hispaniensis-like strain was also assigned to several

ZOTUs (see Table 3.6) and was observed within 93.97% of ticks from all instars. While

Francisella are known tick endosymbionts, this is the first report of F. hispaniensis-like species identified within A. albolimbatum ticks.

A phylogenetic tree of the Francisella-like ZOTUs generated in this study, together with pathogenic and free-living species of Francisella,was made using 431bp 16S gene (Fig

3.14). Despite the relative short fragment length used in this phylogenetic analysis,

Francisella-like ZOTUs generated in this study clustered together with high bootstrap support (86) and were distinct from F. hispaniensis, F. tularensis and Francisella endosymbionts. It is therefore hypothesised that Francisella-like ZOTUs observed within A. albolimbatum ticks represent a novel Francisella, which may be unique to WA.

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Figure 3.13 Phylogenetic analysis of 404bp 16S rRNA gene of Rickettsia using the Neighbor-

Joining algorithm under the assumption of Kimura 2-parameter using a Gamma distribution.

The tree was rooted with members of the order Rickettsiales. Branch lengths indicate the number of character changes and all bootstrap values from 1,000 replications are shown on interior branch nodes.

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Figure 3.14 Phylogenetic analysis of 431bp 16S rRNA gene of Francisella using the

Neighbor-Joining algorithm. The tree was rooted with Piscirickettsia species, the closest relatives to the Francisella. Branch lengths indicate the number of character changes and all bootstrap values from 1,000 replications are shown on interior branch nodes. FLE: Francisella like endosymbiont.

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Two genera belonging to the order Legionellales were identified. ZOTU8 was closely related to a Rickettsiella endosymbiont (KT697666.1; Duron et al. 2016) associated with Ixodes uriae with a 99.3% genetic similarity. This Rickettsiella sp. was observed within 16.38% of ticks from all instars and therefore likely represents a transient Rickettsiella endosymbiont associated with A. albolimbatum ticks.

The second most common ZOTU assigned to the order Legionellales was ZOTU19 with

21,032 sequences in 16 A. albolimbatum ticks (Table 3.5). ZOTU19 had a 100% sequence identity to the known tick-borne pathogen, Coxiella burnetii (LC464975.1). Coxiella sequences generated from NGS were aligned with Coxiella sequences to assist with taxonomic identity. Coxiella sequences obtained (ZOTU19) most closely aligned with Coxiella burnetii and despite the short alignment, the phylogenetic tree exhibited the similar topology with previous studies which have hypothesised the evolution of Coxiella burnetii from a Coxiella endosymbiont within soft ticks (Ornithodoros and Argas spp.) (Fig 3.15) (Duron et al. 2015).

Although 16 A. albolimbatum ticks generated Coxiella burnetii sequences in the NGS assay

(table 3.6), all tick samples were negative for the Coxiella genus-specific PCR assays.

However, the Coxiella-positive control yielded the correct amplicon length for each of the assays and the negative controls did not produce any amplicons.

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Table 3.5 The samples which had reads for ZOTU 19 (Coxiella) shown with their percent relative abundance from each sample and the relevant metadata.

No# Relative Host Sample Reads abundance Instar Status Postcode day Month Year code ZOTU19 (%) R29 10233 33.31 Male Pale 1715 6109 24 January 2015 R28 4343 12.13 Male Pale 1715 6109 24 January 2015 R30 3107 12.51 Nymph Engorged 1715 6109 24 January 2015 R115 1601 16.15 Nymph Engorged 1715 6109 24 January 2015 R7 1402 4.17 Male Fed 1725 6076 14 January 2015 R9 219 6.44 Nymph Engorged 1725 6076 14 January 2015 R32 95 0.19 Nymph Engorged 1717 6556 na na Na R39 8 0.02 Nymph Engorged 1725 6076 14 January 2015 R33 5 0.01 Male Fed 1727 6076 4 January 2015 R3 4 0.02 Male Fed 2743 6068 22 November 2016 R35 3 0.01 Male Fed 2743 6068 22 November 2016 R37 3 0.01 Male Fed 2742 6068 22 November 2016 R5 3 0.01 Male Fed 2743 6068 22 November 2016 R1 2 0 Female Fed 982 6028 7 November 2013 R113 2 0 Female Engorged 1716 6109 24 January 2015 R43 2 0.02 Male Fed 982 6028 7 November 2013

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Table 3.6 Microbial taxa of interest observed in tick samples at a high abundance.

Reads in % Reads in Reads in ZOTU# GenBank # GenBank Identity % Identity negative % in Ticks ticks >999 Ticks controls reads ZOTU1 MK005802.1 Rickettsia endosymbiont of Bemisia tabaci 100.00 1229002 89 77.59 66.38 ZOTU2 KT281843.1 F. hispaniensis strain 2008651S 99.07 915280 63 93.97 65.52 ZOTU3 KT281843.1 F. hispaniensis strain 2008651S 98.14 140185 7 65.52 25.86 ZOTU4 KT281843.1 F. hispaniensis strain 2008651S 98.37 128844 3 75.86 28.45 ZOTU5 KT281843.1 F. hispaniensis strain 2008651S 98.37 130217 11 72.41 18.97 ZOTU6 KT281843.1 F. hispaniensis strain 2008651S 98.60 96089 6 72.41 12.07 ZOTU7 KT281843.1 F. hispaniensis strain 2008651S 98.14 89991 12 22.41 10.34 ZOTU8 KT697666.1 Rickettsiella sp. RKTSLLA_T1705 99.30 60962 14 16.38 6.90 ZOTU9 KT281843.1 F. hispaniensis strain 2008651S 98.60 49163 2 73.28 15.52 ZOTU10 MK304546.1 R. raoultii isolate Tomsk 99.75 47649 0 26.72 1.72 ZOTU11 MK304546.1 R. raoultii isolate Tomsk 99.26 35811 3 20.69 6.90 ZOTU12 KT281843.1 F. hispaniensis strain 2008651S 98.14 28990 0 25.86 7.76 ZOTU13 KT281843.1 F. hispaniensis strain 2008651S 97.90 27271 0 49.14 8.62 ZOTU16 KT281843.1 F. hispaniensis strain 2008651S 98.14 25141 0 54.31 8.62 ZOTU17 KT281843.1 F. hispaniensis strain 2008651S 97.90 21904 2 23.28 6.03 ZOTU19 LC464975.1 C. burnetii CB_30 gene 100.00 21032 0 13.79 4.31 ZOTU29 KT281843.1 F. hispaniensis strain 2008651S 97.90 10989 0 45.69 3.45 ZOTU31 KT281843.1 F. hispaniensis strain 2008651S 97.90 11610 0 35.34 2.59 ZOTU46 KT281843.1 F. hispaniensis strain 2008651S 97.90 4967 0 14.66 0.86 ZOTU57 KT281843.1 F. hispaniensis strain 2008651S 98.14 4130 0 38.79 0.00 ZOTU62 KT281843.1 F. hispaniensis strain 2008651S 97.90 4943 0 32.76 0.00 ZOTU802 MG889594.1 Francisella-like endosymbiont of Dermacentor 96.04 30 0 0.86 0.00 reticulatus ZOTU1048 KY797658.1 Francisella endosymbiont of Haemaphysalis flava 93.71 210 0 5.17 0.00 ZOTU1349 KT281843.1 F. hispaniensis strain 2008651S 95.12 1712 0 61.21 0.00 C. Coxiella; F. Francisella; R. Rickettsia;

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Figure 3.15 Phylogenetic analysis of 431bp 16S rRNA gene of the Coxiella-like species identified within the tick Amblyomma albolimbatum using the Neighbor-Joining algorithm under the assumption of Tamura-3 parameter using a Gamma distribution. The tree was rooted with species of the genus Legionella the close relatives to the genus Coxiella. Branch lengths indicate the number of character changes and all bootstrap values from 1,000 replications are shown on interior branch nodes.

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Chapter 4: Discussion

4.1 TICK IDENTIFICATION

This study morphologically and molecularly identified ticks from the bobtail lizard.

Morphological techniques included the use of microscopy and dichotomous keys, while PCR of the CO1 gene region, coupled with Sanger sequencing, were the designated molecular techniques employed.

4.1.1 Morphological identification

This study morphologically identified 306 Amblyomma ticks collected from the bobtail.

With the exception of 25 ticks, all adult instars were identified as Amblyomma albolimbatum, thus confirming the first hypothesis, that bobtails would be parasitised by one or more of the three main reptile tick species. However, immature tick instars could only be identified to the genus level. As supported by (Sharrad and King, 1981) this is due to the morphological similarities between the immature stages and dearth of reliable morphological characteristics available for Australian ticks.

The most common instar observed was adult male ticks (48%). This finding is supported by Sharrard and Smyth (1981) who also observed a higher relative abundance of males than any other instar of reptile tick species on the lizards and related it to their life traits; i.e. moving around on the host, attaching and detaching, for a period of weeks while waiting for a female to attach (Andrews and Bull, 1980, Sharrad, 1979). The second most common instar was nymphs, followed by adult females, and lastly, only three larvae were identified from this study. This result was unexpected and is not supported by the literature, whereby Sharrad

(1979) and other studies have found a large percentage of larval instars on lizards (Leu et al.,

2010). This incongruence is possibly due to the small size of the larvae and the predilection

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site, which has been reported to be in the ears (Chilton et al., 1992) . Both these factors make larvae difficult to observe and collect, and given the opportunistic method of collecting the samples in this study, may indicate that the collectors were not actively seeking ticks and thus were likely to have missed observations.

4.1.2 Molecular identification

In order to determine the identity of morphologically divergent samples and the identity of nymphs and larvae, a CO1 gene assay was performed(Evans et al., 2019). This assay successfully amplified 100% samples, however only nine samples produced clean chromatograms. This observation was also noted by Evans (2019) who faced challenges amplifying Amblyomma specimens. Unfortunately, despite successful amplification of the

CO1 gene using a nested PCR, described by Song et al (2011), mixed chromatograms in the majority of samples meant they could not be used for further phylogenetic work.

The samples that were successfully sequenced (i.e. clean chromatograms) were all determined to be Amblyomma albolimbatum, resulting in a monophyletic clade with high bootstrap support (Fig 3.6). Within the clade however, there was a high percentage of pairwise distance displayed between the two branches 4.79-5.05% (table 3.3). This is within the species delimitation range for ticks previously described in the literature (Evans et al., 2019; Song et al., 2011). One tick species from the United States, Ixodes scapularis, was previously found to have a 3-7% divergence from the North and South of the Eastern states, however this was over a longer distance than the ticks in this study(Sakamoto et al., 2014). Another example of high genetic divergence, over a considerable geographic distance, was the tick Ixodes eudyptidis. The samples tested had a 4.5–5% divergence from New Zealand to Australia

(Moon et al., 2015).

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The two representative samples used in the current study, were chosen based on their morphological characteristics as well as to encompass a large area to test the hypothesis that a genetic divergence between ticks that are separated by distance can be observed. This hypothesis was supported by the results for nymphs and adults. However, an additional and unexpected finding was that larvae clustered with the southern reference sample, with no pairwise distance (table 3.3), despite being collected more than 200km from the representative sample. This could be due to the translocation of animals from their original location to a wildlife care centre/location (Spencer and Hampton, 2005) or the geographic distance may merely be an artefact of the study; moreover, further samples are required to support the true prevalence of larval ticks on lizards and the population structure in these ticks.

A survey on the electrophoretic variability among populations of six reptile ticks, which used isoenzyme loci to screen A. albolimbatum ticks in Western Australia, observed zero genetic difference in A. albolimbatum from east to west populations spanning over 2000km

(Bull et al., 1984). More recently, however, the barcoding gene, CO1, has been shown to be a more robust marker of speciation (Hebert et al., 2003; Song et al., 2011), revealing a genetic difference of 3% in A. triguttatum specimens in Western Australia (Evans et al., 2019)

One of the limitations of using the CO1 gene for molecular identification, is because it is a mitochondrial gene, it reflects only the evolutionary history of one parent and therefore, has limited applications for assessing recent hybridisation of taxa (Bull et al., 1984; Norval et al., 2019; Sharrad and King, 1981). Hybridisation between the morphologically similar species, A. limbatum and A. alblimbatum has previously hypothesised (Sharrad and King,

1981)Testing this hypothesis would therefore require nuclear gene sequencing and population structure analyses. Recently, Egan and Evans (unpublished data), amplified the nuclear 18S gene of the southern representative sample (R1) and were able to support that R1 was A. albolimbatum. However, as nuclear genes are not appropriate for species level differentiation

(see section 1.2.4), both R1 and the A. albolimbatum sequences from South Australia displayed

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a pairwise difference of 0.01% difference with A. limbatum. Despite this, the 18S distance combined with the high CO1 distance supports that the ticks used in this study are not A. limbatum, which is their morphologically similar sister taxon.

In addition, less than 0.01% difference was reported in the present study between A. albolimbatum and A. vikirri, which is also known to parasitise the bobtail (Keirans et al.,

1996). The CO1 sequence for A. vikirri on GenBank was only 93bp long (accession

#AH011484), in which only 49bp overlapped with the sequences generated from the present study. Thus, no conclusive results were able to be made. However, additional sequencing of

A. vikirri, particularly at the CO1 gene, would help to further delineate the genus Amblyomma, in particular the morphologically similar reptile species of this genus.

4.2 NGS DATA EXPLORATION.

This study is the first to profile the bacterial microbiome communities across all developmental stages of the tick, Amblyomma albolimbatum, which were found on the

Western Australian bobtail. This examination was carried out by targeting the V3–V4 hypervariable regions of the 16S gene with a 2 x 300bp paired-end reads and run on the

Illumina MiSeq NGS platform.

This study demonstrated that tick instar is a significant factor in predicting microbial composition. Both the alpha and beta diversity differed significantly between the tick instars and at both the presence absence and abundance measures it was most significant between males and females. The alpha diversity differed significantly between all life stages, whereby males had the highest diversity in all metrics and nymphs had the second highest (fig 8). This high diversity in males relative to other life stages is supported by previous reports, which have found a significantly high taxonomic diversity in males compared to females(Lalzar et al., 2012; Thapa et al., 2018; Zolnik et al., 2016). This is possibly due to internal hormonal

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changes as previously hypothesised by Zolnik et al. (2016) or competition among symbionts, increased virulence, and bottlenecks experienced by symbionts during vertical transmission

(Bonnet et al., 2017; Lalzar et al., 2012). However, considering the A. albolimbatum ticks’ life cycle, it is probable that male ticks acquire bacteria from the different lizard attachment sites and the lizard’s skin whilst waiting for a mating partner. Furthermore among the ticks bacterial microbiomes there was previously published lizard skin bacteria (Weitzman et al., 2018).

Furthermore, the rarefication plots show that males were not adequately sequenced, so this study may have underestimated the true diversity in males.

The results generated from this study were supported by previous studies on ticks, whereby

Proteobacteria was the most dominant phylum (Egan et al., 2019; Lalzar et al., 2012; Thapa et al., 2018; Zolnik et al., 2016). NGS revealed 3 assigned phyla werepresent, in tick samples, that were at abundances greater than 1%; Proteobacteria (90.3%), Actinobacteria (4.1%),

Firmicutes (3.4%). However, unassigned phyla was present at 1.1%; these were identified as being from the Kingdom Bacteria however were unnasigned at the phyla level. This may indicate novel species, however it is likely they are a sequencing error as the number of ZOTUs were very high for one tick species compared with other studies (Panetta et al., 2017; Zolnik et al., 2016) . However, it was ZOTUs that were used, in this study, to separate closely related bacteria, whereas previous studies used 97% similar OTUs (Edgar, 2016). Ticks analysed in this study harboured one or two, highly abundant family taxa, this is supported by similar evidence overseas that the tick microbiome is dominated by one or two highly abundant endosymbionts (Bonnet et al., 2017; Narasimhan and Fikrig, 2015).

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4.2.1 Francisella

Within the most abundant phylum, Proteobacteria, the most abundant bacterial species observed was Francisella, making up 1,840,933 reads from 16 ZOTUs. Furthermore,

Francisella was the most abundant genus across all tick life stages, with the exception of nymphs. The presence of Francisella was an expected observation, following on from the research of a previous study by Vilcins and colleagues (2009). In this study, despite the use of

Rickettsia primers, Francisella were amplified from A. fimbriatum ticks that were sourced from reptiles in the Northern Territory(Vilcins, 2009). This current study provides further evidence for the Francisella genus in Amblyomma ticks of Australia with A. maculatum being the first Amblyomma species from which Francisella-like DNA was detected (Scoles, 2004).

Furthermore, Francisella has also been reported in high abundance and prevalence of the ornate kangaroo tick, A. triguttatum, in Western Australia removed from red kangaroos (Egan et al., 2019).

Further analyses revealed that the most abundant Francisella sequence (ZOTU #2) had a 99% match to a strain of F. hispaniensis (KT281843). This strain was recently isolated from an acutely febrile woman and identified as the causative agent in the first reported case of

Francisella bacteraemia in Western Australia (Aravena-Román et al., 2015). This patient, who was already immunocompromised, was hospitalised in Perth and the bacterium was isolated from a blood culture. While the transmission of the bacterium in this case was unknown,

Aravena-Román et al. hypothesised that the patient acquired this infection via inhalation of air from a contaminated from air conditioning system. Moreover, this case, along with other literature, provides evidence that immunocompromised patients are particularly vulnerable to infections with uncommon Francisella strains (Huber et al., 2010; Kugeler et al., 2008)(Huber et al, 2010 & Kugeler et al., 2008).

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Francisella is a known tick endosymbiont, forming a necessary component of the ticks’ microbiome to enable the digestion of a blood meal and reproduction, and is also a known zoonotic pathogen (e.g. F. tularensis the causative agent of in the Northern hemisphere)(Bonnet et al., 2017; Foley and Nieto, 2010). At this current time, tick endosymbionts have not been conclusively associated with human illness, however recent literature indicates evidence for challenging this paradigm (Ahantarig et al., 2013; Oskam et al., 2018) Although the abundance of Francisella in the present study was high in bobtail biting ticks, further research is required to determine the importance of this observation; for example, whether this Francisella is capable of horizontal transfer to vertebrates and if the bobtail is a competent reservoir.

One of the first steps to determining the significance of this finding is to identify the reservoir(s) of the Francisella sequenced in the present study. Wildlife and ticks are frequently the reservoirs for tick-borne pathogens and in the Northern hemisphere, many of the tick-borne pathogens’ lifecycles are known(Foley and Nieto, 2010; Levi et al., 2012). A recent report observed the presence of the Northern hemisphere tick-borne pathogen F. tularensis ssp. holarctica from a woman bitten by a wild ringtail possum in Tasmania (Eden et al., n.d.) and although tick and wildlife reservoirs of this bacterium have been restricted to the Northern hemisphere , a recent study reported evidence of an autochthonous infection, thus warranting tick and wildlife screening for a Southern hemisphere for alternate reservoirs of Francisella spp. (i.e. The bobtail) that may cause disease in the human population.

Phylogenetic analysis of the sequences obtained in this study displayed a monophyletic clade of Francisella species with high bootstrap support, displaying a potential novel species and associated strains, however, all branches within the clade displayed low bootstrap support.

Importantly, sequences generated in this study did not cluster with the previously mentioned

F. hispaniensis and the basal lineage of pathogenic species. The 16S rRNA gene is highly

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conserved within the genus and cannot be used to conclusively identify to a novel species and thus, further testing to include additional genes are necessary ((Duron et al., 2017).

.

The ZOTU #2 was also the most abundant Francisella sequence in all life stages and the samples most strongly associated with it were females and larvae, both at 100% prevalence and 64.39% of relative abundance in females and 85.94% in larvae. No statistical difference was found in this ZOTU for fed vs engorged females (p=0.15). Over 90% prevalence was observed in males and nymphs at much lower relative abundance (6% and 22%). The high relative abundance in females and larvae, and between engorged and fed females, points towards a vertical transmission of this Francisella. Furthermore, the overall relative abundance of Francisella is higher in females (68.8%) than males (47.8%). Francisella-like endosymbionts have previously been only associated with the female snake ticks A. veanense and A. helvolum in Thailand (Sumrandee et al., 2014) however, in this study it is found in both adult sexes which is a finding mirrored in A. fimbriatum and A. triguttatum (Egan et al., 2019;

Vilcins, 2009). If all external abiotic and biotic interactions are the same, the difference in the relative amount may be due to internal hormonal mechanisms, which lead to sexual dimorphism, and their specific pheromones that are excreted by the female(Andrews and Bull,

1980). However, the possibility for horizontal transfer of Francisella between bobtail host and tick requires further research.

4.2.2 Rickettsia

Three ZOTUs were identified as Rickettsia, via the database, and Rickettsia was identified as the second most abundant bacteria observed in the ticks in this study. One Rickettsia sequence

(ZOTU1) being the highest overall abundance. Similar to the Francisella sequences, the

Rickettsia sequences showed a sex bias, however, higher bacteria abundance tended towards males (37%) as opposed to females (26%). This male sex bias of Rickettsia has previously been reported in Rhipicephalus turanicus (Lalzar et al., 2012).

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Rickettsia spp. have been reported for a variety of Australian tick species. Rickettsia gravesii has been described from A. triguttatum (Abdad et al., 2017) and Rickettsia sequences have been previously identified in I. tasmani (Vilcins et al., 2009) and the reptile ticks A. fimbriatum (Vilcins, 2009) and B. hydrosauri (Whiley et al., 2016). These Rickettsia are in addition to the known human pathogens (R. australis and R. honei) (Graves and Stenos, 2017).

The two larvae that were profiled in this study exhibited markedly different relative abundance for Rickettsia. The pale larvae only exhibited 0.6%, while the fed larvae presented 10.5% of

ZOTU1, the most abundant identified Rickettsia sequences. Although there were only two larvae used in this study, this result is intriguing because this bacterium may be horizontally transmitted from the reservoir host to the tick as is the case for Lyme borreliosis (Levi et al.,

2012) ) this may provide insight into the tick:host:pathogen lifecycle; further evidence for

Rickettsia being a pathogen as it is low in unfed larvae and adult ticks. Alternatively, it may be important in the moulting success of the tick. This conclusion is supported by the increased abundance in nymphs (41%), and the subsequent decrease in the final life stage, males (37%) and females. This may point towards a possible need for a high amount for moulting success.

This larval abundance may be a false artefact of this study because of such a small number of larval life stages and the bacterium may be undergoing competitive inhibition with other bacteria that the tick has picked up throughout an individual ticks’ life, a theory proposed by

(Zolnik et al., 2016).

The finding of Rickettsia is also important to clinical understanding of the transmission of this potential pathogen. Rickettsia spp. are one of the only currently recognised TBPs affecting people in Australia (Graves and Stenos, 2017). Yet, because of the conserved nature of the

16S gene in Rickettsia, rigorous species delimitation is not possible by the generation of short sequences (Portillo et al., 2017). However, a phylogenetic analysis was undertaken with the

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16S gene sequences, with recognised Rickettsia species. In this analysis the proposed evolutionary species R. bellii sequence was not basal and thus confirmed that the sequences were too short to be appropriately analysed (Stothard et al., 1994). Therefore, it is recommended that a full length 16S gene sequence is generation in addition to other genes.

For example, research into the outer membrane protein A (ompA) gene should be done to determine if any of these Rickettsia are part of the SFG, the group of Rickettsia most strongly associated with reptiles in Australia (Stenos et al., 2003; Vilcins, 2009). These species of

Rickettsia may be of veterinary and medical importance, although these ticks have not been reported to bite humans at this time, documentation of its potential to affect humans should be undertaken.

4.2.3 Order Legionellales

One of the most surprising observations in this study was the evidence of Coxiella burnetii in

13.79% of the ticks tested with NGS (tables 3.5 and 3.6). A sequence belonging to the Coxiella genus was identified that had 100% genetic similarity to C. burnetii, which also phylogenetically clustered with previously sequenced C. burnetii (fig 3.15). Coxiella burnetii is the causative agent of Q fever in people and coxielliosis in livestock and while ticks are important for the natural lifecycle of C. burnetii, they are rarely implicated in the transmission to people (Duron et al., 2015). Q fever is more frequently associated with abattoir workers and people in the veterinary industry who come into contact with contaminated animal secretions and parturient by-products (Oskam et al., 2018). This study’s finding of evidence for the causative agent of Q fever comes with implications for bobtails, wildlife carers and biosecurity agents who work with bobtails and are not immunised against Q fever.

Coxiella burnetii has not been previously associated with reptiles in Australia, with the exception of a recent NGS study on Bothriocroton undatum ticks from goannas, which observed C. burnetii in 1.2% of the reads (Panetta et al., 2017). However, both this study, and

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the aforementioned study failed to amplify C. burnetii via conventional PCR, following the

NGS run. Panetta et al. (2017) used a C. burnetii specific assay that failed in all samples except the positive control. In the present study, a Coxiella genus specific assay was employed to allow for the potential mismatches within the primers. Unfortunately, only the positive control, a Coxiella-like endosymbiont was amplified, previously extracted from the brown dog tick,

Rhipicephalus sanguineus. In an attempt to amplify C. burnetii within A. albolimbatum ticks the following modifications were included: a dilution series was trialled to test for inhibition; and to account for low bacteraemia an increased volume of DNA, increased PCR cycles and extension time were explored.

Although the research laboratory where this experiment took place had previously extracted and amplified Coxiella DNA, the presence of C. burnetii within the A. albolimbatum ticks is unlikely to represent laboratory contamination. This is due to the abundance of C. burnetii reads (> 30%) in one male tick and >10% of the reads observed in one other male and two nymphs all from the same host (table 3.5). Furthermore, C. burnetii was not found in negative controls. The failure to amplify Coxiella may be due to the unanticipated effects of nucleated reptile blood. A future recommendation is that a qPCR should be undertaken to amplify reptile, bacterial and tick DNA in parallel to see the proportions of each in the samples to test if one of these factors is inhibiting the amplification of Coxiella.

Previously C. burnetii agglutinins has been detected and recovered in reptile visceral organs in India (Yadav and Sethi, 1979). There have also been reports of it being associated with the broad host tick A. nuttalli and the monitor lizard tick A. exornatum, both indigenous to Africa

(Burridge, 2001; Nowak-Chmura, 2014). However, a review of this topic found no scientific evidence for the association between reptile ticks and Q fever (Burridge, 2001). This is contrary to the findings in this study, which has identified C. burnetii in reptile ticks, therefore, further investigation is warranted. The results in the present study may be an isolated case,

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nonetheless future work should consider screening bobtails for the presence of C. burnetii.

This is pertinent to wildlife carers and the biosecurity officers as bobtails are frequently traded both legally and illegally within national and international markets, often with ticks still attached (pers. comm. Matt Swan, DBCA).

The bobtail that was associated with ticks with the highest reads for C. burnetii, was further investigated (1715). This was through contact with Kanyana Wildlife Centre (pers. comm

Carol Jackson, Appendix iii). This female bobtail was admitted to the Kanyana Wildlife Centre because of an upper respiratory tract infection and a high tick burden (n = 9) and was pregnant.

This pregnancy is of interest due to the fact that shedding of the C. burnetii is at its highest during birth and coxielliosis can cause abortions in domestic animals (Angelakis and Raoult,

2010). Although information regarding the viviparous bobtail’s offspring could not be acquired, the bobtail reportedly gained sufficient weight to give birth in a healthy manner

(pers. comm. Carol Jackson). Unfortunately, no faecal or blood samples were obtained to determine if the bobtail shed C. burnetii. While it is unknown if the A. albolimbatum ticks acquired C. burnetii via the blood meal from the bobtail host, or from a previous host, this is the first report of C. burnetii associated with A. albolimbatum ticks.

If the A. albolimbatum ticks did acquire C. burnetii via the bloodmeal from the bobtail host, then this may have been due a variety of reasons. The first reason may point to a disquieting new host range for the ornate kangaroo tick, A. triguttatum, which has previously been associated with C. burnetii (Graves and Stenos, 2017). This new host range is hypothesised because of an observation of a female A. triguttatum on a bobtail in South Australia (Petney et al., 2008). This was considered an accidental host because the tick is not normally found in the region and is associated with mammals; it was thought to have been translocated with livestock. However, A. triguttatum is highly prevalent in Western Australia and an infected

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tick may have previously bitten this bobtail, which then transmitted C. burnetii. Lastly, a further reason may be that the omnivorous bobtail fed on infected byproducts.

Another member of the order Legionellales, the genus Rickettsiella, was identified in one

ZOTU (table 3.6). The genus Rickettsiella are known intracellular pathogens of arthropods

(Cordaux et al., 2007) and the Rickettsiella sequence generated in this study matched most closely to a genotype previously identified from the sea bird tick, Ixodes uriae (Duron et al.,

2016). Although the presence of the Rickettsiella in the I. uriae tick was not considered an arthropod pathogen because it was found in all life stages including gravid females, in the present study it was only found in 16.37% of ticks so it may represent a transient pathogenic species. Future studies will be needed to examine the genotype detected here and if it affects tick fitness.

4.2.4 Limitations of NGS

This study amassed a total of 9.7 million raw sequences, and 4.7 million sequences after quality filtering. This amount is fewer than expected according to the Illumina MiSeq protocol for full length V3–V4 reads using a V3 600 cycle kit, which has an expected outcome of 22–25 million sequences. The depth of sequencing is also much less than expected when compared to previous studies targeting this region (Panetta et al., 2017;

Zolnik et al., 2016) Illumina MiSeq, recommends a minimum of 100,000 reads per sample to be sufficient for metagenomic surveys. The V3-V4 16S rRNA hypervariable region is preferred because diversity estimates for bacterial communities are most often the highest(Sperling et al., 2017; Zolnik et al., 2016).

However, there is no consensus as to what should be used for tick microbiome studies.

Furthermore, the alpha rarefication curves did not plateau for the males, however the male instar still showed the highest diversity, so it was considered sufficient for the analyses performed. In future, higher abundance of reads would be recommended, and could be done

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by the use of a blocking primer on Rickettsia or Francisella in order to prevent PCR bias at the preliminary steps of amplification (Gofton et al. 2015).

There is no consensus on what constitutes a contaminant and what is a part of the tick’s microbiota, however the amplification of sequences which aligned with chloroplast and mitochondrial DNA are definite contaminates. They were amplified because both chloroplasts and mitochondria are evolutionarily descended from bacteria, so the 16S genes are nearly homologous between the two (Hanshew et al., 2013). The sterilisation of the tick may not have been stringent enough to remove plant DNA. A future recommendation would be to dissect the tick to determine the organ-specific bacteria. This would also aid in the determination of vertically transmitted bacteria which will be present in the ovaries (Bonnet et al., 2017).

Despite these limitations, this and many studies still provide a valuable source insight into the determination of tick microbiomes (Egan et al., 2019; Panetta et al., 2017). Moreover, although the recommended sequencing depth was not achieved in this study, high quality sequences were generated and were used to assess the bacterial diversity of A. albolimbatum ticks. Lastly, this study showcased novel and previously undescribed taxa of interest.

4.3 CONCLUSIONS

This study is the first to obtain a CO1 gene sequences from morphologically identified

Amblyomma albolimbatum ticks, collected from the Western Australian bobtail, and a relatively high genetic diversity from the Northern and Southern representative ticks was observed. This study provides new molecular data for A. albolimbatum and requires further research to investigate the validity of the Northern and Southern morphotypes identified.

This study achieved its primary aim of next generation sequencing the bacterial microbiome of the tick A. albolimbatum. This is the first study to examine the bacterial microbiome of this tick species across all life stages. The hypothesis that instars have a different bacterial diversity

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to other tick instars was supported by the data whereby the highest alpha diversity was observed in males followed by nymphs.

Proteobacteria was the most abundant phylum across all life stages and genera of significance were identified at high abundance. Taxa of interest included Francisella, Rickettsia, Coxiella and Rickettsiella. Furthermore, for the first time in Australia, this study presents the bacterial communities within reptile ticks and showcases a rich diversity of microbes, including endosymbionts, known tick-associated pathogens and novel microbes which may be putative pathogens. Further studies are needed to determine both the vector competence of A. albolimbatum and the reservoir competence of the bobtails for the bacteria genotypes identified.

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Appendices

Table A1. ZOTUs per tick in above 0, above 30 and number of reads.

RUBY114 7 0 49 Nymph engorged RUBY80 3 0 74 Nymph engorged RUBY54 4 0 87 Male fed RUBY81 3 2 235 Female fed RUBY117 7 2 827 Nymph engorged RUBY83 23 4 2369 Female fed RUBY63 16 7 2799 Male fed RUBY60 27 13 4197 Nymph engorged RUBY18 3 1 4678 Female pale RUBY40 30 4 5126 Nymph engorged RUBY119 11 4 6738 Nymph engorged RUBY24 17 7 7447 Male fed RUBY27 71 25 7653 Nymph pale RUBY115 111 12 9930 Nymph engorged RUBY41 31 7 10944 Nymph engorged RUBY43 38 14 12612 Male Fed RUBY44 17 3 12884 Female Fed RUBY21 26 15 13031 Male Fed RUBY118 29 9 14245 Nymph engorged RUBY25 43 8 14971 Male Fed RUBY23 16 5 15047 Nymph Fed RUBY26 65 51 15228 Nymph Fed RUBY105 32 26 15735 Nymph Fed RUBY17 24 10 17578 Female engorged RUBY66 44 8 18464 Female Fed RUBY77 23 5 19202 Male Fed RUBY99 84 34 19295 Male Fed RUBY92 35 23 19574 Nymph engorged RUBY13 23 7 21362 Male Fed RUBY101 17 3 21437 Larva Fed RUBY113 94 8 21533 Female engorged RUBY2 85 19 21940 Male Fed RUBY19 76 19 22012 Male Pale RUBY85 15 2 22280 Nymph Fed

87

RUBY11 15 2 22334 Female Engorged RUBY35 69 23 22628 Male Fed RUBY15 18 1 23027 Female engorged RUBY45 134 46 23085 Male Fed RUBY70 17 9 23125 Female engorged RUBY16 43 16 23624 Male Pale RUBY57 55 22 24162 Male Fed RUBY6 15 7 24168 Female Fed RUBY49 45 5 24536 Female Fed RUBY3 73 19 24851 Male Fed RUBY34 32 20 25242 Nymph Pale RUBY93 47 25 25637 Male Fed RUBY75 165 55 25720 Male Fed RUBY37 120 13 26139 Male Fed RUBY1 21 6 26158 Female Fed RUBY104 28 16 26766 Nymph engorged RUBY4 43 11 26934 Male Fed RUBY46 87 51 27159 Male Fed RUBY36 19 2 27395 Female Fed RUBY109 28 20 27960 Nymph Fed RUBY86 71 17 29447 Female Fed RUBY97 33 17 29499 Male Fed RUBY90 19 2 30141 Female Fed RUBY14 48 14 30147 Male Fed RUBY103 32 18 30273 Male Fed RUBY29 260 46 30578 Male Pale RUBY76 18 9 30740 Female Fed RUBY107 55 9 31270 Female Fed RUBY48 48 23 31891 Male Fed RUBY102 90 34 32611 Larva Pale RUBY38 23 22 32614 Male Fed RUBY79 42 9 33146 Male Fed RUBY106 37 19 33222 Nymph Fed RUBY30 61 27 33356 Nymph engorged RUBY9 46 19 33407 Nymph engorged RUBY87 147 45 33478 Male Fed RUBY96 20 4 33608 Female Fed RUBY95 18 11 33619 Female Fed RUBY7 64 19 33701 Male Fed RUBY78 36 15 33871 Male Fed RUBY47 310 63 34018 Female Fed RUBY33 42 24 34640 Male Fed RUBY88 31 7 34965 Nymph Fed

88

RUBY39 26 9 35751 Nymph Engorged RUBY28 73 25 35814 Male Pale RUBY89 45 29 35844 Nymph engorged RUBY74 7 1 36795 Nymph engorged RUBY120 61 12 37926 Female Fed RUBY5 54 16 37938 Male Fed RUBY91 6 5 38387 Male Fed RUBY50 31 13 38780 Male Fed RUBY8 19 12 39165 Female engorged RUBY111 111 45 40899 Male Fed RUBY65 64 31 41225 Nymph Fed RUBY67 160 41 41266 Male Fed RUBY72 25 21 41412 Male Fed RUBY82 39 18 41920 Male Fed RUBY51 59 30 42821 Male Fed RUBY64 19 6 42947 Female Fed RUBY100 38 28 45003 Male Fed RUBY84 64 41 46917 Male Fed RUBY61 61 24 47831 Male Fed RUBY98 74 36 48142 Female Fed RUBY73 99 21 48320 Male Fed RUBY56 48 23 48568 Female engorged RUBY10 47 20 48605 Male Fed RUBY32 54 36 49206 Nymph engorged RUBY55 54 23 49394 Male Fed RUBY12 78 16 49901 Male Fed RUBY108 45 18 51950 Nymph fed RUBY42 120 23 53491 Male fed RUBY110 50 37 54018 Nymph engorged RUBY112 82 66 54691 Nymph engorged RUBY52 24 12 56656 Female fed RUBY20 35 9 59497 Female fed RUBY53 96 37 67608 Male fed RUBY22 104 30 71084 Male fed RUBY59 42 22 78978 Male fed RUBY69 29 23 84470 Male fed RUBY94 53 28 86656 Male fed RUBY71 303 130 89144 Male fed RUBY68 48 12 96464 Female fed RUBYIB1 9 1 Control control RUBYNT2 10 10 Control control RUBYEXB3 13 0 Control control RUBYNT1 14 0 Control control

89

RUBYEXB1 28 15 control control RUBYEXB2 41 34 control control

90

Table A.2. Sample metadata used for bioinformatics. #Sample_ID sample-namesample_type DNA_code PI_code Tick_genus Tick_species Tick_genus_sIpNeSciTeAsR status host_commonhost_genus host_species state Suburb post_code day month year Six_season Four_season RUBY1 982_AAF1_R1sample R1 982 Amblyomma albolimbatumA. alb Female fed Bobtail Tiliqua rugosa WA Bussel Hwy x Abbey Drive 6280 7 November 2013 Second SpringSpring RUBY10 1723_AAM1_sRa1m0ple R10 1723 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA West Toodyay 6566 13 December 2014 First SummerSummer RUBY100 1297_AAM1_sRa1m00ple R100 1297 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Baldivis 6171 7 July 2014 Winter Winter RUBY101 1717_AAL1_Rs1a0m1ple R101 1717 Amblyomma albolimbatumA. alb Larva fed Bobtail Tiliqua rugosa WA Chidlow 6556 0 NA 2015 NA NA RUBY102 1717_AAL2_Rs1a0m2ple R102 1717 Amblyomma albolimbatumA. alb Larva pale Bobtail Tiliqua rugosa WA Chidlow 6556 0 NA 2015 NA NA RUBY103 1462_AAM1_sRa1m03ple R103 1462 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Willeton 6155 4 November 2014 Second SpringSpring RUBY104 1319_AAN1_Rsa1m04ple R104 1319 Amblyomma albolimbatumA. alb Nymph engorged Bobtail Tiliqua rugosa WA Gabbadah 6042 10 September 2014 First Spring Spring RUBY105 1755_AAN1_Rsa1m05ple R105 1755 Amblyomma albolimbatumA. alb Nymph fed Bobtail Tiliqua rugosa WA Balga 6061 6 January 2015 First SummerSummer RUBY106 1460_AAN1_Rsa1m06ple R106 1460 Amblyomma albolimbatumA. alb Nymph fed Bobtail Tiliqua rugosa WA Gabbadah 6042 25 September 2014 First Spring Spring RUBY107 1454_AAF1_Rs1a0m7ple R107 1454 Amblyomma albolimbatumA. alb Female fed Bobtail Tiliqua rugosa WA Mt Helena 6082 18 November 2014 Second SpringSpring RUBY108 1736_AAN1_Rsa1m08ple R108 1736 Amblyomma albolimbatumA. alb Nymph fed Bobtail Tiliqua rugosa WA Lesmurdie 6076 20 January 2015 First SummerSummer RUBY109 1740_AAN1_Rsa1m09ple R109 1740 Amblyomma albolimbatumA. alb Nymph fed Bobtail Tiliqua rugosa WA Jandacott 6164 23 December 2014 First SummerSummer RUBY11 2742_AAF1_Rs1a1mple R11 2742 Amblyomma albolimbatumA. alb Female engorged Bobtail Tiliqua rugosa WA Whiteman Park 6068 22 November 2016 Second SpringSpring RUBY110 1738_AAN1_Rsa1m10ple R110 1738 Amblyomma albolimbatumA. alb Nymph engorged Bobtail Tiliqua rugosa WA The Vines 6069 3 January 2015 First SummerSummer RUBY111 1875_AAM1_sRa1m11ple R111 1875 Amblyomma albolimbatumA. alb Male fed Blue-tongue Tiliqua sp. WA Peppemint grove Beach 6271 22 November 2014 Second SpringSpring RUBY112 1751_AAN1_Rsa1m12ple R112 1751 Amblyomma albolimbatumA. alb Nymph engorged Bobtail Tiliqua rugosa WA Jandacott 6164 28 December 2014 First SummerSummer RUBY113 1716_AAF1_Rs1a1m3ple R113 1716 Amblyomma albolimbatumA. alb Female engorged Bobtail Tiliqua rugosa WA Maddington 6109 24 January 2015 First SummerSummer RUBY114 1734_AAN1_Rsa1m14ple R114 1734 Amblyomma albolimbatumA. alb Nymph engorged Bobtail Tiliqua rugosa WA West Toodyay 6566 0 NA 2015 NA NA RUBY115 1715_AAN1_Rsa1m15ple R115 1715 Amblyomma albolimbatumA. alb Nymph engorged Bobtail Tiliqua rugosa WA Maddington 6109 24 January 2015 First SummerSummer RUBY117 1723_AAN1_Rsa1m17ple R117 1723 Amblyomma albolimbatumA. alb Nymph engorged Bobtail Tiliqua rugosa WA West Toodyay 6566 13 December 2014 First SummerSummer RUBY118 1720_AAN1_Rsa1m18ple R118 1720 Amblyomma albolimbatumA. alb Nymph engorged Bobtail Tiliqua rugosa WA Balga 6061 21 January 2015 First SummerSummer RUBY119 1752_AAN1_Rsa1m19ple R119 1752 Amblyomma albolimbatumA. alb Nymph engorged Bobtail Tiliqua rugosa WA West Toodyay 6566 28 December 2014 First SummerSummer RUBY12 2742_AAM3_sRa1m2ple R12 2742 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Whiteman Park 6068 22 November 2016 Second SpringSpring RUBY120 1875_AAF1_Rs1a2m0ple R120 1875 Amblyomma albolimbatumA. alb Female fed Blue-tongue Tiliqua sp. WA Peppemint grove Beach 6271 22 November 2014 Second SpringSpring RUBY13 2843_AAM1_sRa1m3ple R13 2843 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Landsdale 6065 22 October 2017 Second SpringSpring RUBY14 2745_AAM1_sRa1m4ple R14 2745 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Whiteman Park 6068 22 November 2016 Second SpringSpring RUBY15 2744_AAF1_Rs1a5mple R15 2744 Amblyomma albolimbatumA. alb Female engorged Bobtail Tiliqua rugosa WA Whiteman Park 6068 22 November 2016 Second SpringSpring RUBY16 1874_AAM1_sRa1m6ple R16 1874 Amblyomma albolimbatumA. alb Male pale Blue-tongue Tiliqua sp. WA Peppemint grove Beach 6271 3 December 2014 First SummerSummer RUBY17 1874_AAF1_Rs1a7mple R17 1874 Amblyomma albolimbatumA. alb Female engorged Blue-tongue Tiliqua sp. WA Peppemint grove Beach 6271 3 December 2014 First SummerSummer RUBY18 2745_AAF1_Rs1a8mple R18 2745 Amblyomma albolimbatumA. alb Female pale Bobtail Tiliqua rugosa WA Whiteman Park 6068 22 November 2016 Second SpringSpring RUBY19 1872_AAM1_sRa1m9ple R19 1872 Amblyomma albolimbatumA. alb Male pale Blue-tongue Tiliqua sp. WA Peppemint grove Beach 6271 17 November 2014 Second SpringSpring RUBY2 982_AAM1_Rs2ample R2 982 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Bussel Hwy x Abbey Drive 6280 7 November 2013 Second SpringSpring RUBY20 1872_AAF1_Rs2a0mple R20 1872 Amblyomma albolimbatumA. alb Female fed Blue-tongue Tiliqua sp. WA Peppemint grove Beach 6271 17 November 2014 Second SpringSpring RUBY21 1733_AAM1_sRa2m1ple R21 1733 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Wattle grove 6107 1 January 2015 First SummerSummer RUBY22 1732_AAM1_sRa2m2ple R22 1732 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Lesmurdie 6076 4 January 2015 First SummerSummer RUBY23 1732_AAN1_Rsa2m3 ple R23 1732 Amblyomma albolimbatumA. alb Nymph fed Bobtail Tiliqua rugosa WA Lesmurdie 6076 4 January 2015 First SummerSummer RUBY24 1616_AAM1_sRa2m4ple R24 1661 Amblyomma albolimbatumA. alb Male fed Blue-tongue Tiliqua sp. WA Woodridge 6041 4 January 2015 First SummerSummer RUBY25 1718_AAM1_sRa2m5ple R25 1718 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA West Toodyay 6566 11 January 2015 First SummerSummer RUBY26 1718_AAN1_Rsa2m6 ple R26 1718 Amblyomma albolimbatumA. alb Nymph fed Bobtail Tiliqua rugosa WA West Toodyay 6566 11 January 2015 First SummerSummer RUBY27 1718_AAN2_Rsa2m7 ple R27 1718 Amblyomma albolimbatumA. alb Nymph pale Bobtail Tiliqua rugosa WA West Toodyay 6566 11 January 2015 First SummerSummer RUBY28 1715_AAM1_sRa2m8ple R28 1715 Amblyomma albolimbatumA. alb Male pale Bobtail Tiliqua rugosa WA Maddington 6109 24 January 2015 First SummerSummer RUBY29 1715_AAM2_sRa2m9ple R29 1715 Amblyomma albolimbatumA. alb Male pale Bobtail Tiliqua rugosa WA Maddington 6109 24 January 2015 First SummerSummer RUBY3 982_AAM2_Rs3ample R3 982 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Bussel Hwy x Abbey Drive 6280 7 November 2013 Second SpringSpring RUBY30 1715_AAN2_Rsa3m0 ple R30 1715 Amblyomma albolimbatumA. alb Nymph engorged Bobtail Tiliqua rugosa WA Maddington 6109 24 January 2015 First SummerSummer RUBY32 1717_AAN1_Rsa3m2 ple R32 1717 Amblyomma albolimbatumA. alb Nymph engorged Bobtail Tiliqua rugosa WA Chidlow 6556 0 NA 2015 NA NA RUBY33 1727_AAM1_sRa3m3ple R33 1727 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Lesmurdie 6076 4 January 2015 First SummerSummer RUBY34 1727_AAN1_Rsa3m4 ple R34 1727 Amblyomma albolimbatumA. alb Nymph pale Bobtail Tiliqua rugosa WA Lesmurdie 6076 4 January 2015 First SummerSummer RUBY35 2743_AAM1_sRa3m5ple R35 2743 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Whiteman Park 6068 22 November 2016 Second SpringSpring RUBY36 2743_AAF1_Rs3a6mple R36 2743 Amblyomma albolimbatumA. alb Female fed Bobtail Tiliqua rugosa WA Whiteman Park 6068 22 November 2016 Second SpringSpring RUBY37 2742_AAM2_sRa3m7ple R37 2742 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Whiteman Park 6068 22 November 2016 Second SpringSpring RUBY38 1725_AAM2_sRa3m8ple R38 1725 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Lesmurdie 6076 14 January 2015 First SummerSummer RUBY39 1725_AAN2_Rsa3m9 ple R39 1725 Amblyomma albolimbatumA. alb Nymph engorged Bobtail Tiliqua rugosa WA Lesmurdie 6076 14 January 2015 First SummerSummer RUBY4 2743_AAM1_sRa4mple R4 2743 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Whiteman Park 6068 22 November 2016 Second SpringSpring RUBY40 1725_AAN3_Rsa4m0 ple R40 1725 Amblyomma albolimbatumA. alb Nymph engorged Bobtail Tiliqua rugosa WA Lesmurdie 6076 14 January 2015 First SummerSummer RUBY41 1725_AAN4_Rsa4m1 ple R41 1725 Amblyomma albolimbatumA. alb Nymph engorged Bobtail Tiliqua rugosa WA Lesmurdie 6076 14 January 2015 First SummerSummer RUBY42 982_AAM3_Rs4a2mple R42 982 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Bussel Hwy x Abbey Drive 6280 7 November 2013 Second SpringSpring RUBY43 982_AAM4_Rs4a3mple R43 982 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Bussel Hwy x Abbey Drive 6280 7 November 2013 Second SpringSpring RUBY44 982_AAF2_R4s4ample R44 982 Amblyomma albolimbatumA. alb Female fed Bobtail Tiliqua rugosa WA Bussel Hwy x Abbey Drive 6280 7 November 2013 Second SpringSpring RUBY45 1872_AAM2_s4a5mple R45 1872 Amblyomma albolimbatumA. alb Male fed Blue-tongue Tiliqua sp. WA Peppemint grove Beach 6271 17 November 2014 Second SpringSpring RUBY46 1872_AAM3_sRa4m6ple R46 1872 Amblyomma albolimbatumA. alb Male fed Blue-tongue Tiliqua sp. WA Peppemint grove Beach 6271 17 November 2014 Second SpringSpring RUBY47 1872_AAF2_Rs4a7mple R47 1872 Amblyomma albolimbatumA. alb Female fed Blue-tongue Tiliqua sp. WA Peppemint grove Beach 6271 17 November 2014 Second SpringSpring RUBY48 1316_AAM1_sRa4m8ple R48 1316 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Mt Helena 6082 21 September 2014 First Spring Spring RUBY49 679_AAF1_R4s9ample R49 679 Amblyomma albolimbatumA. alb Female fed Bobtail Tiquila rugosa WA Yanchep NP 6035 4 October 2013 Second SpringSpring RUBY5 2743_AAM2_sRa5mple R5 2743 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Whiteman Park 6068 22 November 2016 Second SpringSpring RUBY50 681_AAM1_Rs5a0mple R50 681 Amblyomma albolimbatumA. alb Male fed Bobtail Tiquila rugosa WA Yanchep NP 6035 1 October 2013 Second SpringSpring RUBY51 1317_AAM1_sRa5m1ple R51 1317 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA West Toodyay 6566 11 August 2014 First Spring Winter RUBY52 680_AAF1_R5s2ample R52 680 Amblyomma albolimbatumA. alb Female fed Bobtail Tiquila rugosa WA Yanchep NP 6035 4 October 2013 Second SpringSpring RUBY53 680_AAM1_Rs5a2mple R53 680 Amblyomma albolimbatumA. alb Male fed Bobtail Tiquila rugosa WA Yanchep NP 6035 4 October 2013 Second SpringSpring RUBY54 1325_AAM1_sRa5m4ple R54 1325 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA West Toodyay 6566 10 September 2014 First Spring Spring RUBY55 1295_AAM1_sRa5m5ple R55 1295 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Mt Helena 6082 23 July 2014 Winter Winter RUBY56 983_AAF1_R5s6ample R56 983 Amblyomma albolimbatumA. alb Female engorged Bobtail Tiliqua rugosa WA Bussleton Hospital 6280 5 December 2013 First SummerSummer RUBY57 1296_AAM1_sRa5m7ple R57 1296 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Belmont 6104 7 July 2014 Winter Winter RUBY59 1316_AAM1_sRa5m9ple R59 1316 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Mt Helena 6082 21 September 2014 First Spring Spring RUBY6 1735_AAF1_Rs6ample R6 1735 Amblyomma albolimbatumA. alb Female fed Bobtail Tiliqua rugosa WA Darlington 6070 13 January 2015 First SummerSummer RUBY60 1315_AAN1_Rsa6m0 ple R60 1315 Amblyomma albolimbatumA. alb Nymph engorged Bobtail Tiliqua rugosa WA Huntingdale 6110 20 August 2014 First Spring Winter RUBY61 1459_AAM1_sRa6m1ple R61 1459 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Wattle grove 6107 26 September 2014 First Spring Spring RUBY63 1318_AAM1_sRa6m3ple R63 1318 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA The Vines 6069 27 August 2014 First Spring Winter RUBY64 1298_AAF1_Rs6a4mple R64 1298 Amblyomma albolimbatumA. alb Female fed Bobtail Tiliqua rugosa WA Mt Helena 6082 30 June 2014 Winter Winter RUBY65 1457_AAN1_Rsa6m5 ple R65 1457 Amblyomma albolimbatumA. alb Nymph fed Bobtail Tiliqua rugosa WA Forrestfield 6058 18 November 2014 Second SpringSpring RUBY66 1320_AAF1_Rs6a6mple R66 1320 Amblyomma albolimbatumA. alb Female fed Bobtail Tiliqua rugosa WA Nowergup 6032 5 August 2014 First Spring Winter RUBY67 1320_AAM1_sRa6m7ple R67 1320 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Nowergup 6032 5 August 2014 First Spring Winter RUBY68 1299_AAF1_Rs6a8mple R68 1299 Amblyomma albolimbatumA. alb Female fed Bobtail Tiliqua rugosa WA Lesmurdie 6076 26 June 2014 Winter Winter RUBY69 1458_AAM1_sRa6m9ple R69 1458 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Wallista 6076 18 November 2014 Second SpringSpring RUBY7 1725_AAM1_sRa7mple R7 1725 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Lesmurdie 6076 14 January 2015 First SummerSummer RUBY70 1458_AAF1_Rs7a0mple R70 1458 Amblyomma albolimbatumA. alb Female engorged Bobtail Tiliqua rugosa WA Wallista 6076 18 November 2014 Second SpringSpring RUBY71 1464_AAM1_sRa7m1ple R71 1464 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Lesmurdie 6076 26 September 2014 First Spring Spring RUBY72 1464_AAM2_sRa7m2ple R72 1464 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Lesmurdie 6076 26 September 2014 First Spring Spring RUBY73 1322_AAM1_sRa7m3ple R73 1322 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Chidlow 6556 29 August 2014 First Spring Winter RUBY74 1301_AAN1_Rsa7m4 ple R74 1301 Amblyomma albolimbatumA. alb Nymph engorged Bobtail Tiliqua rugosa WA Mt Helena 6082 23 July 2014 Winter Winter RUBY75 1321_AAM1_sRa7m5ple R75 1321 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA West Toodyay 6566 13 August 2014 First Spring Winter RUBY76 1300_AAF1_Rs7a6mple R76 1300 Amblyomma albolimbatumA. alb Female fed Bobtail Tiliqua rugosa WA Wanneroo 6065 18 July 2014 Winter Winter RUBY77 1300_AAM1_sRa7m7ple R77 1300 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Wanneroo 6065 18 July 2014 Winter Winter RUBY78 3320_AAM1_sRa7m8ple R78 3320 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Jandacott 6164 4 January 2019 First SummerSummer RUBY79 3322_AAM1_sRa7m9ple R79 3322 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Floreat 6014 25 January 2019 First SummerSummer RUBY8 1725_AAF1_Rs8ample R8 1725 Amblyomma albolimbatumA. alb Female engorged Bobtail Tiliqua rugosa WA Lesmurdie 6076 14 January 2015 First SummerSummer RUBY80 3322_AAN1_Rsa8m0 ple R80 3322 Amblyomma albolimbatumA. alb Nymph engorged Bobtail Tiliqua rugosa WA Floreat 6014 25 January 2019 First SummerSummer RUBY81 3319_AAF1_Rs8a1mple R81 3319 Amblyomma albolimbatumA. alb Female fed Bobtail Tiliqua rugosa WA Gosnells 6110 13 December 2019 First SummerSummer RUBY82 3319_AAM1_sRa8m2ple R82 3319 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Gosnells 6110 13 December 2019 First SummerSummer RUBY83 3321_AAF1_Rs8a3mple R83 3321 Amblyomma albolimbatumA. alb Female fed Bobtail Tiliqua rugosa WA Floreat 6014 25 January 2019 First SummerSummer RUBY84 3321_AAM1_sRa8m4ple R84 3321 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Floreat 6014 25 January 2019 First SummerSummer RUBY85 3321_AAN1_Rsa8m5 ple R85 3321 Amblyomma albolimbatumA. alb Nymph fed Bobtail Tiliqua rugosa WA Floreat 6014 25 January 2019 First SummerSummer RUBY86 1872_AAF3_Rs8a6mple R86 1872 Amblyomma albolimbatumA. alb Female fed Blue-tongue Tiliqua sp. WA Peppemint grove Beach 6271 17 November 2014 Second SpringSpring RUBY87 1872_AAM4_sRa8m7ple R87 1872 Amblyomma albolimbatumA. alb Male fed Blue-tongue Tiliqua sp. WA Peppemint grove Beach 6271 17 November 2014 Second SpringSpring RUBY88 1747_AAN1_Rsa8m8 ple R88 1747 Amblyomma albolimbatumA. alb Nymph fed Bobtail Tiliqua rugosa WA The Vines 6069 7 January 2015 First SummerSummer RUBY89 1737_AAN1_Rsa8m9 ple R89 1737 Amblyomma albolimbatumA. alb Nymph engorged Bobtail Tiliqua rugosa WA West Toodyay 6566 28 December 2014 First SummerSummer RUBY9 1725_AAN1_Rsa9mple R9 1725 Amblyomma albolimbatumA. alb Nymph engorged Bobtail Tiliqua rugosa WA Lesmurdie 6076 14 January 2015 First SummerSummer RUBY90 767_AAF1_R9s0ample R90 767 Amblyomma albolimbatumA. alb Female fed Bobtail Tiliqua rugosa WA Mt Helena 6082 15 April 2013 Autumn Autumn RUBY91 767_AAM1_Rs9a1mple R91 767 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Mt Helena 6082 15 April 2013 Autumn Autumn RUBY92 3323_AAN1_Rsa9m2 ple R92 3323 Amblyomma albolimbatumA. alb Nymph engorged Bobtail Tiliqua rugosa WA Floreat 6014 25 January 2019 First SummerSummer RUBY93 3323_AAM1_sRa9m3ple R93 3323 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Floreat 6014 25 January 2019 First SummerSummer RUBY94 3329_AAM1_sRa9m4ple R94 3329 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Southern River 6110 12 March 2019 Second SummAeurtumn RUBY95 1324_AAF1_Rs9a5mple R95 1324 Amblyomma albolimbatumA. alb Female fed Bobtail Tiliqua rugosa WA Wattle grove 6107 19 September 2014 First Spring Spring RUBY96 1744_AAF1_Rs9a6mple R96 1744 Amblyomma albolimbatumA. alb Female fed Bobtail Tiliqua rugosa WA Lesmurdie 6076 16 December 2014 First SummerSummer RUBY97 1744_AAM1_sRa9m7ple R97 1744 Amblyomma albolimbatumA. alb Male fed Bobtail Tiliqua rugosa WA Lesmurdie 6076 16 December 2014 First SummerSummer RUBY98 572_AAF1_R9s8ample R98 572 Amblyomma albolimbatumA. alb Female fed Blue-tongue Tiliqua sp. WA Kilikup 6244 17 October 2013 Second SpringSpring RUBY99 572_AAM1_Rs9a9mple R99 572 Amblyomma albolimbatumA. alb Male fed Blue-tongue Tiliqua sp. WA Kilikup 6244 17 October 2013 Second SpringSpring RUBYEXB1 extraction_blacnokn1trol EB1 extraction_blacnokntrol control control control control control control control control control control control control control control control RUBYEXB2 extraction_blacnokn2trol EB2 extraction_blacnokntrol control control control control control control control control control control control control control control control RUBYEXB3 extraction_blacnokn3trol EB3 extraction_blacnokntrol control control control control control control control control control control control control control control control RUBYIB1 index_blank1control IB1 index_blank control control control control control control control control control control control control control control control control RUBYNT1 no_template_ccoonnttrrooll1 NT1 no_template_ccoonnttrrooll control control control control control control control control control control control control control control control RUBYNT2 no_template_ccoonnttrrooll2 NT2 no_template_ccoonnttrrooll control control control control control control control control control control control control control control control

91

Fig. A.3 Patient information from Kanyana Wildlife centre concerning the bobtail lizard

1715/1716 who was found with C. burnetti positive ticks.

92

Fig A.4 Photos of the female bobtail 715/1716 who was found with C. burnetti positive ticks. Top photo is of the right side of the head and bottom is of the dorsal side of the body.

93

4

post_code

6014 6104 6032 6107 6035 6109 6041 6110 6042 6155 2 6058 6164

] 6271 6061 6171

%

2 .

0 6065 6244

1

/

6068 6271

% 8

. 6069 6280

2 [

6070 6556

2 A

C 6164 6076 6566 C 6082 6109 6244 INSTAR 6070 6280 6104 0 Female 6076 6155 6171 Larva 6556 6107 6082 6042 6058 Male 6032 6035 6065 Nymph 6110 6566 6068 6069 6014 6041

6061

−2 0 2 4 6 8 CCA1 [3.2% / 11.6%]

Fig A.5 Constrained coordinate analysis of postcode and bacterial diversity.

NIL R_NORTH R_SOUTH

Thiotrichales; Francisella 62.3 44.2 32.4 20.5 8.6 84.3 78.1 88.3 49.4 41.4

Rickettsiales; Rickettsia 27.7 38.8 42.4 76.2 89.5 14.7 8.5 5.7 11.5 43.7

Bacillales; Staphylococcus 0 0.1 1.5 0 0 0 4.9 0.1 13.6 3.1

Legionellales; Rickettsiella 0.2 2.9 2.2 0 0 0 0 0 0.1 0

Bacillales; Bacillus 0 2.2 0.1 0 0.1 0 0 0 2.3 0.3

Actinomycetales; Brevibacterium 0 0.3 0.2 0 0 0 2.9 0 9.7 0

p__Unnasigned_Otu212; Unnasigned 3.7 0 0 0 0 0 0 0 0 0

Legionellales; Coxiella 0 1.1 0.9 0 0 0 0 0 0 0.2

Enterobacteriales; Pantoea 2.5 0 0.7 0 0 0 0 0 0 0

Enterobacteriales; Enterobacter 0 1.3 0 0 0 0 0 0 0 0

e e h e e h e a e h

l l l l l l

v

p p p

a a a a a a

r

m m m

a

m M m M m M

y y y

L

e e e

N N N

F F F

Fig A.6 Heatmap of the Northern and Southern A. albolimbatum ticks from CO1 sequencing in comparison to the respective top ten bacterial genera from NGS results.

94

Autumn Spring Summer Winter NA

Proteobacteria; Francisella 99.7 12 77.2 46.7 47.7 42.7 38.9 36.2 46 55.3 1.4 88.3 27.5

Proteobacteria; Rickettsia 0 36.8 11.5 28.4 44.2 43.5 50 39.9 50.8 27.4 75.4 5.7 21.8

Firmicutes; Staphylococcus 0 0 1.9 4.6 0 0 0.2 1.2 0 0.1 0 0.1 8.4

Proteobacteria; Rickettsiella 0 49.1 0.3 0.3 0.1 0 1.4 2.7 0 0 0 0 0

Firmicutes; Bacillus 0 0 0 4.4 0.6 0 0 0 0 0 0 0 0.1

Actinobacteria; Brevibacterium 0 0 1.1 3.3 0 0 0.7 0.2 0 0 0 0 0

Unnasigned; Unnasigned 0.1 0.6 0 0.1 0.7 10.2 1.2 0.2 0 0 0 0 0

Proteobacteria; Uncultured bacterium 0 0.5 0.5 0.6 0.1 0.7 0.6 1.2 1.6 0.6 0 0.2 0.4

Proteobacteria; Coxiella 0 0 0 0 0 0 2.9 1.1 0 0 0 0 0.1

Proteobacteria; Pantoea 0 0 3.4 0 0 0 0 0.8 0 0 0 0 0

e

e

e

e

h

e

e

h

e

e

h

a

h

l

l

l

l

l

l

l

l

v

p

p

p

p

a

a

a

a

a

a

a

a

r

m

m

m

m

a

m

M

m

M

m

M

m

M

y

y

y

y

L

e

e

e

e

N

N

N

N

F

F F F

Fig A.7 Heatmap of the season of sampling of the A.albolimbatum ticks in comparison to the respective top ten bacterial genera from NGS results.

95

Francisellaceae; Francisella 59.4 64 99.9

Rickettsiaceae; Rickettsia 37.7 22.5 0

p__Unnasigned_Otu212; Unnasigned 0 3.9 0

Enterobacteriaceae; Pantoea 0 2.6 0

c__Alphaproteobacteria_Otu81; Uncultured bacter ium 0.8 0.6 0

Planococcaceae; Staphylococcus 0 0.7 0

Bacillaceae; Staphylococcus 0 0.7 0

Brevibacteriaceae; Brevibacterium 0 0.6 0

o__Lactobacillales_Otu58; Enterococcus 0 0.5 0

c__Alphaproteobacteria_Otu1007; c__Alphaproteobacter ia_Otu1007 0.4 0.3 0

Clostridiaceae; Clostridium 0 0.4 0

c__Gammaproteobacteria_Otu482; c__Gammaproteobacter ia_Otu482 0.2 0.2 0

o__Actinomycetales_Otu40; Brevibacterium 0 0.2 0

Coxiellaceae; Rickettsiella 0 0.2 0

o__Sphingomonadales_Otu383; o__Sphingomonadales_Otu383 0.2 0.2 0

Intrasporangiaceae; Arsenicicoccus 0 0.2 0

Moraxellaceae; Acinetobacter 0 0.2 0

o__Actinomycetales_Otu102; Mobilicoccus 0 0.1 0

Micrococcaceae; Arthrobacter 0 0.1 0

c__Alphaproteobacteria_Otu1322; c__Alphaproteobacter ia_Otu1322 0.1 0.1 0

d

d

e

l

e

e

a

f

g

p

r

o

g n e

Fig A.8 Heatmap of Female A. albolimbatum tick status compared to the respective top 20 bacterial genera from the NGS results.

96

Female Nymph

Proteobacteria; Francisella

Proteobacteria; Rickettsia

Unnasigned; Unnasigned

Firmicutes; Staphylococcus

% Read Abundance Proteobacteria; Pantoea 75

50

25

Proteobacteria; Rickettsiella

Proteobacteria; Uncultured bacterium

Proteobacteria; Sphingomonas

Proteobacteria; f__Sphingomonadaceae_Otu286

Proteobacteria; Halomonas

d d le d d le e fe a e fe a rg p rg p o o g g n n e e

Fig A.9 Heatmap of the status of female and nymph A. albolimatum ticks compared against relative abundance of the top ten genera of bacteria from NGS results.

97

MDS ordination

0.4 ]

% INSTAR

6 .

5 Female

1 [

0.0

Male

2

. s

i Nymph

x A

−0.4

−1.0 −0.5 0.0 0.5 Axis.1 [26.3%]

Fig A.9 MDS plot showing the differences in beta-diversity between tick instars at ZOTU taxa abundance data based on Bray-Curtis dissimilarity.

98